Nucleic Acids Research

Weusten, Jos J. A. M.;Carpay, Wim M.;Oosterlaken, Tom A. M.;van Zuijlen, Martien C. A.;van de Wiel, Paul A.

doi: 10.1093/nar/30.6.e26pmid: 11884645

Abstract For quantitative NASBA-based viral load assays using homogeneous detection with molecular beacons, such as the NucliSens EasyQ HIV-1 assay, a quantitation algorithm is required. During the amplification process there is a constant growth in the concentration of amplicons to which the beacon can bind while generating a fluorescence signal. The overall fluorescence curve contains kinetic information on both amplicon formation and beacon binding, but only the former is relevant for quantitation. In the current paper, mathematical modeling of the relevant processes is used to develop an equation describing the fluorescence curve as a function of the amplification time and the relevant kinetic parameters. This equation allows reconstruction of RNA formation, which is characterized by an exponential increase in concentrations as long as the primer concentrations are not rate limiting and by linear growth over time after the primer pool is depleted. During the linear growth phase, the actual quantitation is based on assessing the amplicon formation rate from the viral RNA relative to that from a fixed amount of calibrator RNA. The quantitation procedure has been successfully applied in the NucliSens EasyQ HIV-1 assay. Received September 24, 2001; Revised and Accepted January 28, 2002. INTRODUCTION In recent years, HIV-1 viral load measurement has become a generally applied tool for anti-retroviral therapy monitoring and evaluation of clinical trials. Three main application areas have been identified. First, a baseline level of HIV-1 predicts efficacy of treatment, second, the level of viral load drop upon treatment is an important predictor for the durability of the treatment and, third, the absolute value of HIV-1 RNA upon treatment has prognostic significance (1–4). Commercially available quantitative HIV-1 viral load assays are based on NASBA, bDNA and RT–PCR (5–7). We have developed a system based on combining NASBA amplification and real-time detection utilizing molecular beacons in order to meet increasing needs with respect to analytical assay performance as well as user convenience and high throughput. Molecular beacons are hairpin-shaped oligonucleotides (Fig. 1). The stem of the beacon is formed by complementary sequences at both ends of the oligonucleotide. A fluorescent label and a quenching group are attached at the two ends of the molecule. The stem holds these two groups in close proximity to each other, causing the fluorescence of the fluorophore to be quenched by energy transfer. When the molecular beacon encounters a target molecule containing a sequence that is complementary to the loop sequence, a hybrid is formed. This hybridization forces the stem apart and causes the fluorophore and the quencher to move away from each other. Now a fluorescence signal can be generated that is not quenched (see for example 8). In recent papers (9,10) we reported the development of a quantitative HIV-1 viral load assay based on NASBA using molecular beacons that hybridize to single-stranded amplicons. The NASBA-driven RNA growth and the subsequent beacon binding yield a fluorescence signal that reflects the events occurring in the reaction tube. As such, detection is performed during the amplification and not afterwards, as is done in former NASBA-based assays (11,12). Analogous to these former assay systems, a fixed amount of calibrator RNA is added to the sample to serve as an internal quantitation standard. This calibrator RNA differs from the sample HIV-1 RNA by only a small part of the sequence, which enables specific hybridization with a sequence-specific molecular beacon. In the assay two different molecular beacons are used, each with a different loop structure to ensure specific binding to either the HIV-1 sample RNA or calibrator amplicons, and each labeled with a different fluorophore to allow specific detection. These two molecular beacons are present at the start of the NASBA reaction and as both the endogenous sample RNA and the calibrator RNA are amplified, two fluorescence signals are produced concomitantly with amplification. Based on these observed fluorescence profiles, the amount of HIV-1 RNA in the original sample is quantified. We have developed a mathematical tool to achieve this goal, which is described below. QUALITATIVE DESCRIPTIONS OF THE PROCESSES Molecular biology of the NASBA process The generally accepted scheme describing the NASBA process is presented in Figure 2 (12–14). The process makes use of two primers, which are short single-stranded DNA fragments, and three enzymes. The viral RNA strands that are present in the original sample are indicated as being the sense strand. Primer P1, after binding to the RNA, is elongated by reverse transcriptase (AMV-RT) to yield a DNA:RNA hybrid, the RNA strand of which is hydrolyzed by RNase H. Then primer P2 can bind, which is elongated by AMV-RT activity to yield a double-stranded DNA molecule. Primer P1 is designed such that it contains a nucleotide sequence that, once it is double-stranded, forms a T7 RNA polymerase promoter site so that antisense RNA copies can be generated using the DNA template. From this RNA new copies of DNA are formed in a process similar to that described for the sense strand, but now P2 is the first primer to bind. The final product is a DNA species with a double-stranded promoter region which is sufficient to allow T7 RNA polymerase to use it as a template, so generating more copies of the antisense RNA. Although the reaction sequences starting from the sense and antisense RNA strands form biochemically somewhat different forms of DNA (Fig. 2), they are considered to be kinetically identical and are both referred to as copy DNA (cDNA). As long as the concentrations of the primers are not rate limiting, the formation of antisense RNA proceeds exponentially with time (see below), although quantitatively important amounts are not yet formed due to the efficiency of the cDNA-producing but RNA-consuming machinery. Initially, the concentration of primer relative to the total concentration of amplifiable RNA is very high. When assaying a high viral load sample, the number of RNA copies entering amplification is of the order of 105–106, whereas the number of P1 and P2 oligonucleotides is 1012. Therefore, the concentration of primer is not rate limiting and relatively small amounts of primer are consumed in depletion of the initially present pool of sense RNA copies. At some time point, obviously, the primer concentrations do become rate limiting, and decline to practically zero. At this time point the cDNA levels have reached their maximum and antisense RNA production proceeds at high speed, so this transition time interval is short. From now on the only reaction that can proceed is T7 RNA polymerase-mediated formation of antisense RNA from the cDNA templates. As will be shown below, accumulation of RNA is linear with time and quantitatively important amounts of RNA are actually produced. This phase is referred to as the transcriptional phase and lasts for a considerable amount of time. Typically, more than 1014 RNA molecules can be generated in a NASBA reaction. If amplification is allowed to proceed over a long enough time period, the RNA formation rate starts to decrease due to depletion of the nucleotide pool and due to (thermal) degradation of the enzyme activity. For the current assay system, however, this moment is so late that there is no need to include it in the models. Principles of quantitation During the amplification process two types of RNA are present: the endogenous viral HIV-1 RNA (WT RNA) and a calibrator RNA (Q RNA) that is added to the sample in a fixed amount prior to extraction of the RNA from the sample. Both types of RNA are converted into cDNA, so that there is competition between the RNA types for the primer pool, and the relative amounts of RNA that were originally present in the sample determine the relative amounts of cDNA formed. As the amount of cDNA formed determines the RNA production rate in the transcriptional phase, the relative rates of RNA formation in this phase directly reflect the relative concentrations of WT and Q RNA in the sample. As the concentration of Q RNA is known, the WT level can be computed. The antisense RNA molecules themselves are not detectable. Therefore, molecular beacons are added that bind to the RNA amplicons. This binding takes time. The shape of the fluorescence curve that is obtained depends on two processes: the NASBA-driven time-dependent growth in RNA levels and binding of the beacon to this RNA. In order to allow quantitation, both processes are modeled, yielding an equation for the fluorescence curve. The transcription rates are derived from this curve. A number of reference samples of which the HIV-1 RNA content is known are subjected to the assay to assess the relation between the transcription rates ratio on the one hand and the WT RNA concentration in the sample on the other. MATHEMATICAL MODELING In this part of the paper the relevant processes will be modeled. For the quantitation procedure the combined early exponential and late linear growth in RNA levels is crucial, so these properties will be dealt with first. Then the development of the fluorescence signal over time will be discussed. NASBA-driven RNA growth To model the major kinetic properties of the amplification process as depicted in Figure 2, a set of reaction rate equations has to be derived. As noted above, the initial concentrations of primers is at least 106 times higher than the total concentration of amplifiable RNA, so during the first phases of amplification they can be considered to be constant and not rate limiting. If in addition (pseudo) first order kinetics is assumed for each reaction, a set of linear differential equations is obtained. In the derivations below, the individual nucleic acid species are represented by abbreviations, with the prefix s for sense, a for antisense and c for copy. The individual reaction rate constants are represented by the symbols c1–c11 and the brackets [ ] indicate concentrations. The set of differential equations can be written in matrix notation as equation S1 with the matrix A defined according to S2 As noted, the primer concentrations [P1] and [P2] are considered to be constant in this phase of the amplification, hence their appearance in the matrix A. In general, the solution of such a set of linear first order differential equations can be written as νiexp(ξit), with ξi the eigenvalues of matrix A and νi reflecting eigenvectors. It can be shown that exactly one of the eigenvalues is positive and that the others are either real negative numbers or complex numbers with a negative real part. This latter is due to the fact that the T7 polymerase catalyzed reaction, reflected by c6, is considerably slower than the primer binding reactions reflected by c7[P2] and c10[P1]. It follows, after an initialization phase, that the single positive exponent term remains and growth is essentially exponential. When the primer pool is depleted, the reaction rate equations become much simpler. The only terms remaining for antisense RNA growth are d[aRNA]/dt = c6[cDNA] 2 Since at this time point the concentration of cDNA is constant, it follows that the increase in antisense RNA levels is linear with time. Obviously, between the phases described by equations 1 and 2 there is a transition phase in which the primer concentrations cannot be considered to be constant, as they decrease to zero. The set of equations 1 is no longer linear and the equations cannot be solved in general anymore. For the current modeling studies this is of minor importance; the distinction of the exponential and linear growth phases is the relevant issue. The time point of primer depletion marks the transition of these two growth phases. Beacon binding to RNA: the central differential equation Quantitation in the assay is based on detection of the molecular beacon signal that is generated after formation of the RNA:beacon hybrid (Fig. 1). This binding can be described by the simple chemical reaction scheme: k2 R + B Z RB k1 with R the antisense RNA species, B the molecular beacon and RB the RNA:beacon complex. The symbols k1 and k2 are the two reaction rate constants that describe the association and dissociation rates. As the beacon binding reaction follows second order kinetics (8), the system can be described by the differential equation d[RB(t)]/dt = k1[R(t)][B(t)] – k2[RB(t)] 3 with the brackets [ ] indicating concentrations, t the time and (t) indicating time dependency. The design of the beacon is such that binding of the beacon to RNA proceeds rapidly, whereas dissociation proceeds relatively slowly. In addition, the RNA levels increase rapidly with time. Therefore, the contribution of the dissociation reaction can be ignored and k2 can be set to zero. The total concentration of beacon is constant and is referred to as Btot. These two considerations can be applied to differential equation 3, yielding d[RB(t)]/dt = k1[R(t)]{Btot – [RB(t)]}. This is a simple differential equation, which can be solved in general. It follows that [RB(t)] = Btot – Iexp{–k1∫[R(t)]dt} 4 with I an integration constant, to be defined later. The fluorescence signal The total fluorescence signal is determined by the concentration of the RNA:beacon complex and the amount of signal generated by this hybrid. In addition, the free beacon contributes a signal and there is (potentially) some background signal. Therefore, the total fluorescence signal is given by Y(t) = ε0 + ε1[B(t)] + ε2[RB(t)] = ε0 + ε1{Btot – [RB(t)]} + ε2[RB(t)], 5 with Y(t) the fluorescence signal as a function of time t, ε0 the background level signal and ε1 and ε2 the coefficients giving the amount of signal for the free and RNA-bound beacons. Note that it follows that ε1 < ε2 as the fluorescence signal increases after opening of the beacon. Combining equations 4 and 5 yields Y(t) = ε0 + ε2Btot –(ε2 – ε1)Iexp{–k1∫[R(t)]dt}. 6 An equation to describe NASBA-driven RNA growth It is clear that equation 6 is not practically useful as long as no mathematically manageable equation can be given for the RNA levels [R(t)]. There is no need to include all individual reaction rate constants of equation 1 in this RNA growth describing equation, but it is of importance that this equation combines an initial exponential growth phase with a later linear growth phase. As the fluorescence levels start to plateau long before the nucleotide pool becomes rate limiting, there is no need to include a late plateau phase of the RNA levels in the equations. An equation that combines an early exponential with a late linear growth and that is integratable as well is [R(t)] = α1α2[eα2(t – α3)/(1 + eα2(t – α3))]ln(1 + eα2(t – α3)), 7 with α1, α2 and α3 the parameters that describe the growth (all positive). Equation 7 can be approximated by [R(t)] = α1α2e2α2(t – α3) for α2(t – α3) < –5.1 (exponential growth) and by [R(t)] = α1α22(t – α3) for α2(t – α3) > +4.4 (linear growth) with errors <1%. It follows that the transcription rate is given by α1α22. From equation 7 it follows that ∫[R(t)]dt = α1[ln(1 + eα2(t – α3))]2. 8 No integration constant is needed here as it can be incorporated in the constant I of equation 4. Overall equations Combining equations 6 and 8 yields an equation that describes the development of the fluorescence signal over time: Y(t) = λY0 – (λ – 1)Y0exp{–k1α1[ln(1 + eα2(t – α3))]2}, 9 with λY0 = ε0 + ε2Btot and (λ – 1)Y0 = (ε2 – ε1)I, hence characterizing the integration constant I from equation 4. Equation 9 describes a fluorescence curve with horizontal asymptotes given by Y(t) = Y0 and Y(t) = λY0. The exact shape of the curve is defined by the values of α1 and α2, whereas α3 defines the location of the curve on the time axis. Note that in equations 7–9 the time appears only relative to α3; the exact moment of the start of the reaction is not part of the equations. Note that the parameters k1 and α1 make their appearance as the product k1α1 only. An example of a fluorescence curve and the corresponding RNA levels according to equations 7 and 9 is presented in Figure 3. Note that the fluorescence levels reach a plateau after ∼50 min due to exhaustion of the beacon pool, whereas the RNA levels still increase linearly with time. Interpretation of the α3 parameter The parameter referred to as α3 has a clear physical interpretation. Using Matlab Simulink a simulation tool was developed (not shown) to quantitatively simulate the processes involved in the NASBA reaction. This simulation tool confirmed the initial exponential and later linear growth in RNA levels and revealed that the overall RNA curves could be described adequately by equation 7. In addition, it showed that the value of α3 is a good approximation of the time that the primer pool is depleted. Therefore, it is referred to as the ‘time to primer depletion’. Since WT and Q RNA make use of the same primer pool, it will be clear that a single value of α3 is used to describe both the WT and calibrator fluorescence curves. THE QUANTITATION PROCEDURE Determining the values of the kinetic parameters in an individual amplification result During an amplification reaction with molecular beacons, the fluorescence signals of the WT and calibrator RNA are measured continuously. Both the time and the observed signals are monitored. When data collection is complete, the WT and calibrator curves are fitted using non-linear regression methods using equation 9. The two fluorescence curves are fitted simultaneously, as they share a common value of α3. In total nine different parameter values are fitted: Y0, λ, k1α1 and α2 for the WT curve, Y0, λ, k1α1 and α2 for the calibrator curve and the common value of α3. After the fit has been completed, the RNA growth curve according to equation 7 can be reconstructed. Assessing the value of the quantitation variable Now that the values of the relevant kinetic parameters have been assessed and the RNA growth curve reconstructed, the formation rate of RNA in the transcriptional phase can be computed. The ratio of the WT over Q RNA transcription rates defines the quantitation variable. From equation 7 it follows that the transcription rate is given by α1α22 (see also Fig. 3). As the values of k1 and α1 cannot be estimated separately (equation 9), it follows that the quantitation variable is defined as the k1α1α22 ratio. Quantitation From equation 2 it follows that the ratio of the transcription rates (i.e. the α1α22 ratio) depends on the ratio of the cDNA levels formed from both species of RNA (endogenous virus RNA and calibrator RNA). The ratio of the cDNA levels depends directly on the ratio of the concentrations entering amplification. As long as the ratio of the values of the k1 parameters for the two species of RNA is constant for a given batch, it follows that there is a direct relation between the RNA input ratio and the k1α1α22 ratio. To assess the exact relation with the concentration ratio in the original sample, a series of samples with known input levels and a fixed concentration of calibrator was subjected to the assay. The logarithm of the observed transcription rate ratio is plotted against the logarithm of the nominal input and a straight line is fitted through the data. The slope and intercept so obtained are used as batch parameters and are used for quantitation of unknown samples. The exact values depend on batch-dependent reagent properties, like the exact concentration of calibrator that is used, the quality of the enzymes, the properties of the molecular beacons (such as the value of the k1 parameter ratio) and others. MATERIALS AND METHODS To study the kinetics of RNA and DNA formation, the NASBA procedure as described elsewhere was used (11,12). The kinetics were studied using a well-defined RNA species used as calibrator Qa in the NucliSens HIV-1 QT assay, a non-homogeneous NASBA-based assay using end-point detection based on electrochemiluminescence (ECL). This calibrator RNA was subjected to amplification directly, without using any sample pretreatment steps like extraction. Using specific ECL probes, the total amount of antisense RNA and sense DNA formed can be studied. Amplification was performed under standard conditions (11,12) and was terminated after the required time period by placing the tubes on ice. For the current study the exact amounts formed are of minor importance, only the time dependency being relevant. The results are expressed as ECL counts. Experiments using the molecular beacon-based assay were performed as described elsewhere (10,15). RESULTS Kinetics of NASBA-driven DNA and RNA growth Figure 4 shows the observed ECL signals reflecting DNA and RNA growth. Note that the results confirm the predicted linear RNA growth after longer time periods. A straight line is fitted through the linear part, which crosses the time axis at ∼23 min. According to the principles described above, this is approximately the time point of primer depletion (α3). RNA production in the early phase is quantitatively too low to allow confirmation of the exponential growth phase. The DNA levels reach a plateau. Note that the observed value of α3 based on RNA growth is in line with the start of the plateau of the DNA levels. Beacon-based assay properties An example of a set of fluorescence curves as obtained in a single amplification reaction using endogenous WT RNA and calibrator is presented in Figure 5. The figure is similar to Figure 3, but now real data are presented for both fluorescence curves simultaneously. The fluorescence curves were fitted to the model of equation 9 and the RNA growth curves were reconstructed using the parameter values so obtained with equation 7. Note that the parameter fitted is k1α1, whereas it is α1 that makes its appearance in the RNA growth equation (equation 7). Therefore, the reconstructed RNA growth curves as presented are defined up to some multiplication factor. The figure shows the linear increase in RNA levels with time. The slope of this part of the RNA growth curve forms the basis of quantitation. To study the relation between the logarithm of the estimated transcription rate ratio (the quantitation variable) and the logarithm of the input level, a dilution series of an in vitro cultured HIV-1 viral stock solution, in which the number of particles was quantitated by electron microscopy (11,16), was subjected to the beacon-based assay at two different test sites. A typical example is shown in Figure 6. A linear relation is obtained between the logarithm of the input and the quantitation variable value. The slope and intercept of this line are assessed using straightforward linear regression techniques and are used as batch parameters. In Figure 6 the dose–response relations as obtained using a single batch at two different sites are presented. DISCUSSION In the current paper modeling has been described that forms the basis of quantitation procedures that can be used in a viral load assay using NASBA as the amplification technique and homogeneous detection using molecular beacons. A mathematical description of the processes involved yielded an equation that describes the dynamics of fluorescence signal development with time as a function of several kinetic parameters. The modeling is complicated by the fact that all processes proceed simultaneously. Kinetic measurements are often performed by monitoring the rate of (to take an example in this field) binding of beacon to a fixed amount of target RNA. The very nature of the NASBA system is of course that the RNA levels are not constant with time, so their levels should be modeled as well. The overall fluorescence curve is the combined result of NASBA-driven RNA growth and binding of the beacon to these continuously changing RNA levels. A number of approximations and assumptions were required to obtain a manageable equation describing the fluorescence. For example, Figure 2 represents a model of the complete NASBA scheme and in equation 1 all individual reactions are assumed to follow first order kinetics. In the light of this, an RNA growth curve (equation 7) could be designed that combines the major properties of early exponential and late linear growth with mathematical manageability (reflected by equation 8). Nevertheless, in spite of the approximations and assumptions, overall an equation was derived that gives an adequate description of the observed fluorescence curve. Also, it was found to provide a good basis for quantitation. Figure 4 shows, amongst others, that the RNA levels are still increasing after 90 min amplification and that deviation from linear growth is not yet detectable. The amplification time used in the beacon-based NucliSens assay is only 60 min. This supports the claim that it is of no use to include in the equations the late plateau in RNA levels due to depletion of the nucleotide pool. The RNA growth curve as given by equation 7 does not include such a plateau. The results shown in Figure 6 indicate that the dose–response relation is linear over at least 3.5 log. In addition, it shows that the lines are identical for both study sites, indicating that reader-to-reader differences and similar sources of variation play no role of any relevance. A single fluorescence curve is described by five parameters, while for the simultaneous fit procedure nine parameters are used. Although this is quite a large number, it is clear that it cannot be reduced. After all, a curve is defined by the level of its start signal, its final plateau signal, the time point of the start of fluorescence growth and the steepness of the fluorescence curve. This already yields four parameters for a single curve. Then the shape of the lower bend is different from that of the upper bend and they bear no relation to one another as the molecular biological processes responsible for the two bends are fundamentally different. This introduces a fifth parameter. As indicated, time makes its appearance in the equations in terms of t – α3 only. This is important from an experimental point of view. Due to this, it is not required to monitor the time between the start of the amplification (mixing of the components) and the start of the fluorescence measurements. Obviously, it is necessary to have a value of α3 large enough to allow estimation of the Y0 parameter values. Note that the parameters k1 and α1 make their appearance in the final fluorescence describing equation only as the product k1α1. As a result, the quantitation variable is defined as the k1α1α22 ratio, although the k1 value has nothing whatsoever to do with the transcription rate. As long as the ratio of the k1 values for the endogenous WT RNA and the calibrator RNA can be considered to be constant, it does not matter due to the use of the batch parameters. Apart from the NASBA-based system, other amplification-based systems using real-time detection are being developed making use of PCR technology (for example 17–21). Quantitation using (RT–)PCR and molecular beacons can be based on the so-called ‘cycle threshold’ CT, which is the number of cycles required to obtain a measurable signal (19–21). Due to the fundamentally different nature of the processes in NASBA, where all reactions proceed simultaneously and continuously, a different quantitation strategy is required. In addition, in the NASBA-based real-time assay a calibrator is used throughout the procedure to monitor procedural losses and to monitor the effects of unknown sample factors that might interfere with the amplification kinetics. In conclusion, the major processes in the NASBA process and homogeneous detection using molecular beacons could be modeled and incorporated in a manageable mathematical model. This model was found to provide a good basis for quantification of endogenous WT RNA. * To whom correspondence should be addressed. Tel: +31 411 654 509; Fax: +31 411 654 311; Email: [email protected] View largeDownload slide Figure 1. Mode of action of a molecular beacon. In the closed form (left) the fluorophore F and quencher Q are in close proximity, preventing generation of a signal. After binding to amplicon RNA (right) the beacon opens and a signal can be generated that is not quenched. View largeDownload slide Figure 1. Mode of action of a molecular beacon. In the closed form (left) the fluorophore F and quencher Q are in close proximity, preventing generation of a signal. After binding to amplicon RNA (right) the beacon opens and a signal can be generated that is not quenched. View largeDownload slide Figure 2. Schematic overview of the NASBA process. View largeDownload slide Figure 2. Schematic overview of the NASBA process. View largeDownload slide Figure 3. Theoretical example of a fluorescence curve with corresponding RNA growth curve according to equations 7 and 9. Some relevant quantities are indicated. View largeDownload slide Figure 3. Theoretical example of a fluorescence curve with corresponding RNA growth curve according to equations 7 and 9. Some relevant quantities are indicated. View large Download slide View large Download slide Figure 4. Observed levels of RNA (upper) and DNA (lower) during amplification. The amount of product formed was determined by measuring the ECL signal (arbitrary units). The figure shows the linear RNA growth and the timing of the plateau in DNA levels. View large Download slide View large Download slide Figure 4. Observed levels of RNA (upper) and DNA (lower) during amplification. The amount of product formed was determined by measuring the ECL signal (arbitrary units). The figure shows the linear RNA growth and the timing of the plateau in DNA levels. View largeDownload slide Figure 5. Example of a test result: two fluorescence curves and reconstructed RNA growth curves. The individual fluorescence measurements are indicated by circles, the fitted fluorescence curves by solid lines and the reconstructed RNA growth curves by dashed lines. The vertical reference line indicates the value of α3. View largeDownload slide Figure 5. Example of a test result: two fluorescence curves and reconstructed RNA growth curves. The individual fluorescence measurements are indicated by circles, the fitted fluorescence curves by solid lines and the reconstructed RNA growth curves by dashed lines. The vertical reference line indicates the value of α3. View largeDownload slide Figure 6. The relation between the nominal HIV-1 RNA input tested and the observed logarithm of the k1α1α22 ratio (the quantitation variable) as obtained by assaying dilution series of a viral stock at two different sites with one batch. The results (means ± SD) are shifted somewhat to the left or right to increase readability. View largeDownload slide Figure 6. The relation between the nominal HIV-1 RNA input tested and the observed logarithm of the k1α1α22 ratio (the quantitation variable) as obtained by assaying dilution series of a viral stock at two different sites with one batch. 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Bucka, Alexander;Stasiak, Andrzej

doi: 10.1093/nar/30.6.e24pmid: 11884643

Abstract In recent years there has been growing interest in the question of how the particular topology of polymeric chains affects their overall dimensions and physical behavior. The majority of relevant studies are based on numerical simulation methods or analytical treatment; however, both these approaches depend on various assumptions and simplifications. Experimental verification is clearly needed but was hampered by practical difficulties in obtaining preparative amounts of knotted or catenated polymers with predefined topology and precisely set chain length. We introduce here an efficient method of production of various single-stranded DNA knots and catenanes that have the same global chain length. We also characterize electrophoretic migration of the produced single-stranded DNA knots and catenanes with increasing complexity. Received December 14, 2001; Revised and Accepted January 24, 2002. INTRODUCTION Single-stranded DNA is an ideal nano-engineering material (1,2). Regions of poly(dT) [in the absence of their complement, poly(dA)] are very flexible (3), while regions that are prepared to anneal with each other form a well-defined right-handed, anti-parallel helix with ∼10 bp per turn (4). These two elements, flexible linkers and right-handed helical blocks, permit the design of a great variety of knots and catenanes. Figure 1 illustrates our experimental design for forming different knots and catenanes. The design of catenanes is conceptually simple. We use two different 60mers and a short ‘band-aid’ oligonucleotide. With the exception of regions that can be used for the pairing of two 60mers and the region of complementarity to the band-aid, all the remaining nucleotides are dTs. The band-aid function is to aid one of the 60mers to have its ends brought together for subsequent ligation, while the second 60mer uses the first 60mer as a band-aid (Fig. 1, left). The length of the pairing region between the two 60mers determines the extent of catenation. We have used complementary regions of 12, 20 and 30 bp hoping to obtain singly, doubly and triply linked catenanes, respectively. Upon ligation with T4 DNA ligase, the band-aid is removed during gel electrophoresis performed under DNA-denaturing conditions. The design of knots is conceptually more complex and is a modification of the method originally proposed by Seeman and co‐workers (5,6). A 120mer oligonucleotide contains short complementary sequences forming stem and loop regions (Fig. 1, right). Stem-forming regions are both not-divided so that upon annealing they can form relatively stable duplex regions. One of the loop-forming sequences is, however, divided between two ends of the 120mer and therefore the overall stability of the loop region is decreased as compared with the stem region. In addition, formation of the loop is a quasi-third-order type of reaction requiring the simultaneous interaction of three quasi-independent sequence elements, while the stem formation, which is a quasi-second-order type of reaction, requires the interaction of two sequence elements. This sequence design favors a folding process in which the stem should form first and is followed by pairing in the loop. Ligation in the loop region (Fig. 1, right) leads then to the formation of knots. Notice that loop formation preceding formation of the stem would result in the formation of ‘unknots’ (unknotted circles). By varying the length of stem and loop one can direct formation of different knots. We have used stems and loops of 11 or 15 bp. MATERIALS AND METHODS Oligonucleotides were obtained from Medprobe, Oslo. T4 DNA ligase and exonuclease I, exonuclease III and T4 polynucleotide kinase were supplied by New England Biolabs. Fine chemicals were purchased from Roche and Fluka. The oligonucleotides of 60 or 120 bases, but not the short band-aids or ‘stem splitters’, were phosphorylated by T4 polynucleotide kinase. For the synthesis of knots and catenanes, phosphorylated oligonucleotides and, where appropriate, band-aids or stem splitters were mixed, heated to 80°C for 5 min, annealed at 60°C for 30 min and subsequently allowed to cool to room temperature. The mixtures were ligated overnight at 4°C, at a concentration of 0.4 pmol/ml DNA, with 25 U/ml T4 DNA ligase. If needed, linear molecules left after the ligation were removed by treatment with a mixture of two exonucleases (Escherichia coli exonuclease I and exonuclease III; New England BioLabs and Fermentas, respectively). For the formation of catenanes we used the following oligonucleotides (written in the 5′→3′ direction): ‘first circle’ (Fig. 1, left), GCTCC(T)10GTGTCCTGTCCTGTCCTCTCGTCGTCCCTC(T)10CCTGC; the ‘band-aid’ closing the first circle, AAGGAGCGCAGGAA; the second circle with 12 nt complementary to the first one, GACAGG(T)48CGAGAG; the second circle with 20 nt complementary to the first one, GACAGGACAG(T)40ACGACGAGAG; and the second circle with 30 nt complementary to the first circle, GACAGGACAGGACAC(T)30GAGGGACGACCAGAG. For the formation of knots we have used the following 120mers: constructs with stem and loop regions of 11 bp each (see Fig. 4A), GAGGAG(T)24GTCCTGTCCTG(T)14CTCCTCGTCCC(T)14CAGGACAGGAC(T)24GGGAC; constructs with stem and loop regions of 11 and 15 bp, respectively (see Fig. 4C), GAGGAGAG(T)22GTCCTGTCCTG(T)12CTCTCCTCGTCCCTC(T)12CAGGACAGGAC(T)22GAGGGAC; and constructs with stem and loop regions of 15 bp each (see Fig. 4C), GAGGAGAG(T)20GTGTCCTGTCCTGTC(T)10CTCTCCTCGTCCCTC(T)10GACAGGACAGGACAC(T)20GAGGGAC. Stem-splitting oligonucleotides were complementary to one of the regions forming a stem (see Fig. 3B). To block stem formation in the case of 120mers with a stem of 15 bp we used the following 24mer: AAAAGTGTCCTGTCCTGTCAAAA. The ligation products were separated by denaturing polyacrylamide gel electrophoresis [12% polyacrylamide (PAA), 1× TBE, 8 M urea at 25 V/cm for 2 h] or by alkaline agarose gel electrophoresis [4% NuSieve Agarose (FMC Bioproducts), 50 mM NaOH, 1 mM EDTA at 4 V/cm for 6 h], followed by staining with SYBR Gold (Molecular Probes) and visualization on a UV transilluminator. RESULTS AND DISCUSSION Formation of catenanes Ligation reactions generating different catenanes and knots were monitored by denaturing polyacrylamide gel electrophoresis. To distinguish between contaminating linear molecules like unligated substrates or ligated linear multimers, digestion with exonucleases was performed. Figure 2A shows analysis of the ligation reaction with substrates for the generation of catenanes. In the case of the 12 nt-long pairing region between the two 60mers we observed the generation of three major species resisting digestion by exonucleases (Fig. 2A, lane 2). Quickly migrating bands consist of monomeric circular 60mers, as demonstrated by the fact that a band at this position is also obtained when the reaction includes only one of the 60mers and its 14 nt-long band-aid (data not shown). The two slowly migrating species are presumably singly and doubly linked catenanes. DNA species with similar electrophoretic mobilities were also produced in reactions where the pairing regions between two 60mers were 20- and 30-nt long (Fig. 2A lanes 3 and 4, and 5 and 6, respectively). However, each of these reactions produced only one major slowly migrating band consisting presumably of doubly and triply linked catenanes, respectively. We have verified by restriction digestion that these species indeed consist of catenated monomeric circles and not of dimeric rings that could also be produced in these ligation reactions and would then have a similar electrophoretic mobility to catenated circles. The respective DNA species were gel purified and annealed with a band-aid oligonucleotide to create a recognition site for HhaI restriction endonuclease. Upon partial digestion of catenanes composed of two 60mers, equal intensity bands of linear and circular 60mers should appear since only one of the composing circles could be cleaved by HhaI. Figure 2B shows that this is indeed the case. It should be mentioned here that putative dimers composed of two 60mers that are complementary to the band-aid oligonucleotide should produce upon partial digestion linear 120mers and linear 60mers but not circular 60mers. Dimers composed of 60mers that are not complementary to the band-aid oligonucleotide should be resistant to HhaI restriction endonuclease. Mixed dimers are unlikely to form as there is no band-aid for their ligation, but even if they were formed their digestion cannot produce linear and circular 60mers. Electrophoretic migration of catenanes One of our experimental expectations was to observe ‘natural’ electrophoretic separation of singly, doubly and triply linked catenanes according to their extent of catenation (7). However, under applied electrophoretic conditions (12% PAA, 1× TBE, 8 M urea at 25 V/cm for 2 h) this was apparently not the case. Figure 2A shows, for example, that the catenanes expected to be triply linked migrated at a speed that situated them between catenanes expected to be singly and doubly linked. It is possible, however, that 8 M urea is not sufficient to permanently melt duplex regions, especially when these regions are kept in proximity by catenation and when gels are run at an ambient temperature. In the absence of complete melting, the electrophoretic migration would not only reflect different topology of catenated molecules but would be additionally affected by the different ratio of single- to double-stranded DNA regions in different constructs. To be sure that duplex regions in catenanes are completely melted we decided to analyze gel-purified individual species by electrophoresis on alkaline agarose gels. Figure 2C shows that under such conditions the constructs engineered to produce triply linked catenanes indeed migrate distinctly quicker than constructs intended to produce doubly linked catenanes. Out of the two species produced using 12 bp complementarity between two 60mers, one migrated as expected for singly linked catenanes while the second one co-migrated with a construct expected to produce doubly linked catenanes. Therefore, the migration of completely melted constructs on alkaline agarose gels strongly suggests that our constructs are indeed singly, doubly and triply linked catenanes as predicted in our construction schemes. Figure 2D shows schematically how two different catenanes can be formed in the case of 12 nt complementarity between the two single-stranded DNA oligonucleotides. Upon annealing the two strands are kept together but the nick provides a freedom of rotation. Single-stranded DNA flanking the duplex region is not constrained and this causes a small energy difference between arrangements that upon ligation creates singly linked catenanes (Fig. 2D, left) or doubly linked catenanes (Fig. 2D, right). In the case of constructs with 20 and 30 nt complementarity, the single-stranded DNA flanking the duplex region is already constrained and therefore energy-minimizing topoisomers are produced in the ligation reaction. Probably the mixture of two topoisomers could be obtained by ligation of constructs with 25–26 nt-long complementarity. Formation of knots Figure 3A shows an analysis of the ligation reaction with 120mers designed to form knots and having their stem and loop regions set to 11 bp each (see Fig. 4A). Lane 2 (after the marker lane containing 120 and 240 nt-long linear single-stranded oligonucleotides) analyzes the whole ligation reaction and lane 3 contains the species that resisted digestion by exonucleases. Lane 4 analyzes the ligation reaction in the presence of the oligonucleotide that forms a stable duplex with one of the regions required for stem formation. This oligonucleotide should specifically inhibit the formation of knots while not interfering with efficient ligation in the loop region. The presence of the stem-splitting oligonucleotide leads, therefore, to the formation of unknotted circles (Fig. 3B). Lane 5 shows the effect of exonuclease treatment on DNA species present after the ligation performed in the presence of the stem-splitting oligonucleotides and reveals the position of circular 120mer. Comparison of lanes 2, 3, 4 and 5 clearly indicates that in the absence of the stem splitter two types of knots are produced when stem and loop regions were 11 bp each. Notice that upon elimination of linear molecules, knots are almost exclusive products of the ligation reaction and that unknotted circles are hardly ever formed (except when promoted by the stem splitter). Lanes 6 and 7 analyze the ligation reaction with 120mers having their loop and stem regions set to 11 and 15 bp, respectively. Exonuclease treatment (lane 7) reveals that with this construct only one type of knot is formed. This type of knot migrates like the ‘quicker knot’ produced using 120mers with the stem and loop having 11 bp each. Co-migration of these two knotted species is best seen in samples not treated with exonucleases (lanes 2 and 6) and therefore containing small amounts of linear dimers produced during ligations (240 nt long). These linear dimers serve as good internal markers demonstrating that the quicker knot in lane 2 and the only knot in lane 6 show practically the same electrophoretic mobility and therefore are presumably of the same type. Figure 3C shows a comparison of ligation reactions involving 120mers that had their stems and loops set to three different combinations of lengths. All reaction products were, in addition, treated with exonucleases to eliminate linear species. Pairs of corresponding lanes analyze the ligation reactions performed without and with stem splitters. It can be seen again that in the presence of stem splitters ligation reactions are directed to the formation of circular products. In the absence of stem splitters, however, one observes the highly specific formation of knots where the knots seem to get more complex as the length of stem and loop regions increases. Several earlier studies have demonstrated that in the case of knotted DNA molecules of the same length but forming different knots, the more complex knots migrate quicker than simpler knots (8–12). As in Figure 3A, we observe here (Fig. 3C) that 120mers having their loop and stem regions both set to 11 bp produce two different knots (lane 2). The quicker, and therefore more complex, one seems to be identical (with correction for the gel ‘smiling’) to the only knot produced by 120mers with stem and loop regions of 11 and 15 bp, respectively (lane 4). The 120mers having their loop and stem regions both set to 15 bp produce three different knots, the major one shows much higher electrophoretic migration and is presumably significantly more complex than the other two. The slowest knot, which is produced in very small quantities, is most likely of the same type as the one produced by 120mers with stem and loop regions of 11 and 15 bp long, respectively. The third knot is also produced in small amounts and it migrates very closely to the slowest knot suggesting that these two knots have the same minimal number of crossings (11,12). Electrophoretic migration of knots and catenanes As already mentioned, it is not certain whether, in PAA gels containing urea but run at an ambient temperature, all duplex regions of DNA are permanently melted, especially when these regions are kept in close proximity by the knotting of short oligonucleotide strands. In the absence of complete melting, the electrophoretic migration would not only be a function of different topologies of knotted molecules but would additionally be affected by the different ratio of single- to double-stranded regions in different constructs. To be sure that duplex regions in knots are completely melted we decided to use harsher denaturing conditions such as in alkaline agarose gels. We gel purified three principal types of knots obtained with different 120mers and analyzed them together with three principal catenanes obtained with pairs of 60mers. Figure 3D (lanes 1–3) reveals an ‘arithmetic progression’ of electrophoretic migration of the catenanes that are most likely singly, doubly and triply linked and, thus, have two, four and six crossings in their standard representation. The three principal knotted species (Fig. 3C, lanes 4–6) also show an ‘arithmetic progression’ of electrophoretic migration with the progression steps practically identical to those observed in catenated species (lanes 1–3). Since the analyzed knots and catenanes have the same overall molecular mass the similarity of the progression steps suggests that the analyzed knots progressively increase their minimal crossing number in steps of two. From the design of three different oligonucleotides used for knotting we expected to obtain knots with three, five and seven crossings (Fig. 4). Therefore, the observed differences in electrophoretic migration of produced knots are consistent with our knotting strategy. Identification of different knots formed with the same single-stranded DNA constructs Figure 4 shows the ‘construction plans’ used to generate various knots using oligonucleotides with stem and loop regions of specific length. The figure also shows the expected principal types of knots in their minimal crossing representation. The notation used, 31, 52 and 74, is from the standard tables of knots (13–15), where the main number corresponds to the minimal number of crossings of a given knot and the index number indicates the tabular position of this knot among the knots with a given minimal number of crossings. Presetting stem and loop regions to 11 bp each, we expected efficient formation of trefoil knots having a mathematical notation 31. The stem of 11 bp and the loop of 15 bp were expected to promote the formation of a knot with five crossings known as 52 knot. Oligonucleotides with stem and loop both set to 15 bp were expected to ligate forming knots with seven crossings, which have the mathematical notation 74. The gel analysis of major products is consistent with our expectations, but only in the case of the oligonucleotide with the stem of 11 bp and the loop of 15 bp did the ligation result in formation of unique type of knot. Why were other constructs not so specific? Figure 5A and B shows schematically how an oligonucleotide with stem and loop regions of 11 bp each can form 31 and 52 knots in almost equal proportions. The 11 bp complementarity in the loop can lead, during annealing, to the creation of two principal situations. In the first case there are two right-handed crossings in the loop and the formed knot is the trefoil. In the second case there are three right-handed crossings in the loop and the formed knot is 52. When the complementarity in the loop is of 15 nt the three crossings are almost always created and, therefore, the oligonucleotide with this sequence setting produces almost exclusively 52 knots (see Fig. 3). Figure 5C–E illustrates how an oligonucleotide with stem and loop regions preset to 15 bp each can form two different five crossing knots in addition to 74 knots (Fig. 5C). A 15 bp stem may be at the allowable size limit for this short oligonucleotide and would require high stretching of poly(T) regions (5,6). Such a stretching would oppose completion of the last turn in the stem and this would result then in the formation of 52 knots (Fig. 5D). In a small fraction of the molecules the stress may result in a stem (Fig. 5E). In such a situation the free ends have to cross-back in order to anneal in an anti-parallel way with their complementary sequence. Such a cross-back would lead to the formation of 51 knots (Fig. 5E). The analysis of possible knotting outcomes (Figs 4 and 5) and the observed pattern of electrophoretic migration of formed knots strongly suggests that we have correctly recognized the formed knots. The length of single-stranded DNA (120 nt) was too small to apply the RecA-coating method that was used previously to determine the formed knot types by electron microscopy (16). RecA filaments have the persistence length of 630 nm (17) and therefore could not accommodate the curvature required to follow the knots of the total length of ∼80 nm. We would like to stress here that the presented method allows very efficient creation of different knots and catenanes of the same total chain size. The knotting scheme we propose results in almost 100% efficiency of knot formation. Some of the tested substrates produce specifically only one type of knot or catenane while other substrates can result in the formation of two or more species of knots and catenanes. Out of a given type of knot only one ‘enantiomer’ is obtained and these knots with sequence-specific modifications could be used for ‘templating’ chirality specific synthesis reactions. We observed here that, in the denaturated form, single-stranded knots and catenanes of the same total length react very similarly to the increase of minimal crossing number. An increase of crossing number by two causes a similar increase of electrophoretic migration in knots and catenanes. However, knots seem to be shifted in relation to catenanes and the trefoil knot with three crossings migrates like the catenane with four crossings. Likewise, the knot with five crossings migrates like a catenane with six crossings. This result suggests that for the same number of minimal crossings short single-stranded knots are less compact than the catenanes of the same total length. Earlier studies analyzing electrophoretic migration of double-stranded knots and catenanes of the same total length showed that for the same minimal number of crossings knots and catenanes show essentially the same electrophoretic migration (18). However, in these earlier studies, DNA molecules were 6.4 kb long and this meant that their trajectory was essentially not affected by inter-segmental repulsion and would follow random walk trajectory restricted to a given topological space. In our case the dimensions of molecules are of the same order as the Debye length at applied conditions (19) and therefore their trajectory would be non-random and would rather tend to minimize the self-repulsing energy (20). ACKNOWLEDGEMENTS We thank Jacques Dubochet for his constant support and interest in the project. This work was supported by grants from the Swiss National Science Foundation (31-61636.00 and 31-58841.99) and from the Human Frontiers Science Program. * To whom correspondence should be addressed. Tel: +41 216 924282; Fax: +41 216 924105; Email: [email protected] Present address:Alexander Bucka, European Patent Office, Directorate 1.2.12, NL-2280 HV Rijswijk, The Netherlands View largeDownload slide Figure 1. General strategy to form catenanes and knots. (Left) Sequence of events leading to the formation of catenanes. Three different oligonucleotides were used in each ligation reaction. The two pairing and ligation events are commutative, i.e. their relative order is not important for the final outcome (formation of catenanes). (Right) Sequence of events leading to formation of knots. Only one type of oligonucleotide is used in the ligation reaction. The sequence of pairing in the stem and loop region is crucial for the final outcome. Knots are only formed when stem is formed first. Unknotted circles are formed when the pairing in the loop precedes this in the stem. View largeDownload slide Figure 1. General strategy to form catenanes and knots. (Left) Sequence of events leading to the formation of catenanes. Three different oligonucleotides were used in each ligation reaction. The two pairing and ligation events are commutative, i.e. their relative order is not important for the final outcome (formation of catenanes). (Right) Sequence of events leading to formation of knots. Only one type of oligonucleotide is used in the ligation reaction. The sequence of pairing in the stem and loop region is crucial for the final outcome. Knots are only formed when stem is formed first. Unknotted circles are formed when the pairing in the loop precedes this in the stem. View largeDownload slide Figure 2. Gel-electrophoretic analysis of ligation reactions leading to the formation of various catenanes. (A) Exonuclease eliminates linear single-stranded DNA molecules and reveals DNA species that are likely to be catenanes. Notice that 12 nt-long complementary sequences between the two 60mers (annealing of these sequences is required for the circularization of the ‘second’ 60mer, but can also lead to formation of linear multimers composed of these 60mers) result in frequent formation of dimeric and trimeric linear species, which are therefore exonuclease sensitive. Longer complementary sequences eliminate the formation of these dimeric and trimeric linear molecules but promote the formation of species with high molecular weight (see the top of the gel). (B) Partial restriction cleavage of one of the 60mers unambiguously demonstrates the catenated nature of DNA species that we expected to be catenated. In lanes 2 and 4 notice the equal intensity of circular and linear 60mers released from gel-purified slowly migrating species. In lane 6 circular 60mers are partially cut by HhaI and get partially converted to linear 60mers. (C) In alkaline agarose gels catenanes separate according to their expected linking number. M, marker lane containing linear 116mer. (D) Formation of singly and doubly linked catenanes in the case of 12 nt-long complementarity between the two rings. With these substrates the catenanes with linking number 1 (left) and 2 (right) show small energy difference and therefore form with similar probability. (A and B) PAA-denaturing gels with urea and (C) alkaline agarose NuSieve gel 4%. View largeDownload slide Figure 2. Gel-electrophoretic analysis of ligation reactions leading to the formation of various catenanes. (A) Exonuclease eliminates linear single-stranded DNA molecules and reveals DNA species that are likely to be catenanes. Notice that 12 nt-long complementary sequences between the two 60mers (annealing of these sequences is required for the circularization of the ‘second’ 60mer, but can also lead to formation of linear multimers composed of these 60mers) result in frequent formation of dimeric and trimeric linear species, which are therefore exonuclease sensitive. Longer complementary sequences eliminate the formation of these dimeric and trimeric linear molecules but promote the formation of species with high molecular weight (see the top of the gel). (B) Partial restriction cleavage of one of the 60mers unambiguously demonstrates the catenated nature of DNA species that we expected to be catenated. In lanes 2 and 4 notice the equal intensity of circular and linear 60mers released from gel-purified slowly migrating species. In lane 6 circular 60mers are partially cut by HhaI and get partially converted to linear 60mers. (C) In alkaline agarose gels catenanes separate according to their expected linking number. M, marker lane containing linear 116mer. (D) Formation of singly and doubly linked catenanes in the case of 12 nt-long complementarity between the two rings. With these substrates the catenanes with linking number 1 (left) and 2 (right) show small energy difference and therefore form with similar probability. (A and B) PAA-denaturing gels with urea and (C) alkaline agarose NuSieve gel 4%. View largeDownload slide Figure 3. Gel-electrophoretic analysis of various knots produced upon ligation of knot-forming oligonucleotides. (A) Exonuclease digestion and elimination of knots by the presence of stem splitter allows us to identify knotted DNAs. (B) The principle action of stem splitter. (C) Knots with increasing electrophoretic mobility are formed as pairing regions in stem and loop get longer. (D) Comparison of migration of knots and catenanes with the same total size but with different complexity. (A and C) PAA-denaturing gels with urea and (D) alkaline agarose NuSieve gel 4%. View largeDownload slide Figure 3. Gel-electrophoretic analysis of various knots produced upon ligation of knot-forming oligonucleotides. (A) Exonuclease digestion and elimination of knots by the presence of stem splitter allows us to identify knotted DNAs. (B) The principle action of stem splitter. (C) Knots with increasing electrophoretic mobility are formed as pairing regions in stem and loop get longer. (D) Comparison of migration of knots and catenanes with the same total size but with different complexity. (A and C) PAA-denaturing gels with urea and (D) alkaline agarose NuSieve gel 4%. View largeDownload slide Figure 4. Design of various knots: (A) stem and loop of 11 bp each, (B) stem and loop of 11 and 15 bp, respectively, and (C) stem and loop of 15 bp each. Notice that each pairing region is right-handed and that regions of pairing are anti-parallel. This constraint limits the possible outcomes of the reactions but with oligonucleotides 11/11 and 15/15 we also observed other types of knots (see Fig. 5). It may not be obvious how the ‘constructs’ drawn on the left can be manipulated into standard diagrams of corresponding knots. Arrows show which type of movement of the colored portion needs to be performed to obtain standard representations of knots. View largeDownload slide Figure 4. Design of various knots: (A) stem and loop of 11 bp each, (B) stem and loop of 11 and 15 bp, respectively, and (C) stem and loop of 15 bp each. Notice that each pairing region is right-handed and that regions of pairing are anti-parallel. This constraint limits the possible outcomes of the reactions but with oligonucleotides 11/11 and 15/15 we also observed other types of knots (see Fig. 5). It may not be obvious how the ‘constructs’ drawn on the left can be manipulated into standard diagrams of corresponding knots. Arrows show which type of movement of the colored portion needs to be performed to obtain standard representations of knots. View largeDownload slide Figure 5. Formation of two or three different knots from the same substrate. (A and B) Notice that pairing in the loop region extends in both cases over 11 bp and that in both cases the regular right-handed helix is formed in the loop region. The resulting knots are therefore almost equally likely to be formed and, in fact, that was what we observed (see Fig. 3A and C). (C–E) Notice that (C), (D) and (E) differ only in ‘phasing’ of crossings in the 15 bp long stem. In (C) pairing starts after the first crossing, in (D) a base or two pair before the first crossing and in (E) four or five bases pair before the first crossing. These differences determine how the out-coming ends exit the stem to pair within the loop. Notice that in (E) the out-coming ends have to cross-back as otherwise their polarity is not compatible with pairing in the loop. In (A–D) arrows show which type of movement of the colored portion needs to be performed to obtain standard representations of knots. In the case of (E) we invite readers to take a string or rubber tubing and perform the transition from the left to right and contrary. View largeDownload slide Figure 5. Formation of two or three different knots from the same substrate. (A and B) Notice that pairing in the loop region extends in both cases over 11 bp and that in both cases the regular right-handed helix is formed in the loop region. The resulting knots are therefore almost equally likely to be formed and, in fact, that was what we observed (see Fig. 3A and C). (C–E) Notice that (C), (D) and (E) differ only in ‘phasing’ of crossings in the 15 bp long stem. In (C) pairing starts after the first crossing, in (D) a base or two pair before the first crossing and in (E) four or five bases pair before the first crossing. These differences determine how the out-coming ends exit the stem to pair within the loop. Notice that in (E) the out-coming ends have to cross-back as otherwise their polarity is not compatible with pairing in the loop. In (A–D) arrows show which type of movement of the colored portion needs to be performed to obtain standard representations of knots. In the case of (E) we invite readers to take a string or rubber tubing and perform the transition from the left to right and contrary. References 1. Winfree,E., Furong,L., Wenzler,L.A. and Seeman,N.C. ( 1998) Design and self-assembly of two-dimensional DNA crystals. Nature , 394, 539–544. Google Scholar 2. Braun,E., Eichen,Y., Sivan,U. and Ben-Yoseph,G. ( 1998) DNA-templated assembly and electrode attachement of a conducting silver wire. Nature , 391, 775–778. Google Scholar 3. 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Biol. Chem. , 260, 4975–4983. Google Scholar 9. Stark,W.M., Grindley,N.D., Hatfull,G.F. and Boocock,M.R. ( 1989) Resolvase-catalysed reactions between res sites differeing in the central dinucleotide of subsite I. EMBO J. , 10, 3541–3548. Google Scholar 10. Bates,A.D. and Maxwell,A. ( 1993) In Rickwood,D. and Male,D. (eds), DNA Topology. IRL Press, Oxford. Google Scholar 11. Crisona,N.J., Kanaar,R., Gonzalez,T.N., Zechiedrich,E.L., Klippel,A. and Cozzarelli,N.R. ( 1994) Processive recombination by wild-type Gin and an enhancer-independent mutant. Insight into the mechanisms of recombination selectivity and strand exchange. J. Mol. Biol. , 243, 437–457. Google Scholar 12. Stasiak,A., Katritch,V., Bednar,J., Michoud,D. and Dubochet,J. ( 1996) Electrophoretic mobility of DNA knots. Nature , 384, 122. Google Scholar 13. Alexander,J.W. and Briggs,G.B. ( 1927) On types of knotted curves. Ann. Math. , 28, 562–586. Google Scholar 14. Rolfsen,D. ( 1976) Knots and Links. Publish or Perish Press, Berkeley, CA. Google Scholar 15. Adams,C.C. ( 1994) The Knot Book. W.H. Freeman and Company, New York, NY. Google Scholar 16. Krasnow,M.A., Stasiak,A., Spengler,S.J., Dean,F., Koller,T. and Cozzarelli,N.R. ( 1983) Determination of the absolute handedness of knots and catenanes of DNA. Nature , 304, 559–560. Google Scholar 17. Egelman,E.H. and Stasiak,A. ( 1986) Structure of helical RecA-DNA complexes. Complexes formed in the presence of ATP-gamma-S or ATP. J. Mol. Biol. , 191, 677–698. Google Scholar 18. Wasserman,S.A., Dungan,J.M. and Cozzarelli,N.R. ( 1985) Discovery of a predicted DNA knot substantiates a model for site-specific recombination. Science , 229, 171–174. Google Scholar 19. Eisenberg,D. and Crothers,D. ( 1979) Physical Chemistry with Applications to the Life Sciences .  The Benjamin/Cummings Publishing Company, Inc., Menlo Park, CA. Google Scholar 20. O’Hara,J. ( 1998) Energy of knots. In Stasiak,A., Katritch,V. and Kauffman,L.H. 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Gueldener, U.;Heinisch, J.;Koehler, G. J.;Voss, D.;Hegemann, J. H.

doi: 10.1093/nar/30.6.e23pmid: 11884642

Abstract Heterologous markers are important tools required for the molecular dissection of gene function in many organisms, including Saccharomyces cerevisiae. Moreover, the presence of gene families and isoenzymes often makes it necessary to delete more than one gene. We recently introduced a new and efficient gene disruption cassette for repeated use in budding yeast, which combines the heterologous dominant kanr resistance marker with a Cre/loxP-mediated marker removal procedure. Here we describe an additional set of four completely heterologous loxP-flanked marker cassettes carrying the genes URA3 and LEU2 from Kluyveromyces lactis, his5+ from Schizosaccharomyces pombe and the dominant resistance marker bler from the bacterial transposon Tn5, which confers resistance to the antibiotic phleomycin. All five loxP–marker gene–loxP gene disruption cassettes can be generated using the same pair of oligonucleotides and all can be used for gene disruption with high efficiency. For marker rescue we have created three additional Cre expression vectors carrying HIS3, TRP1 or bler as the yeast selection marker. The set of disruption cassettes and Cre expression plasmids described here represents a significant further development of the marker rescue system, which is ideally suited to functional analysis of the yeast genome. Received November 15, 2001; Revised and Accepted January 21, 2002. INTRODUCTION In 1996 the complete sequence of the genome of the budding yeast Saccharomyces cerevisiae was published (1) and functional analysis of the more than 6000 genes found in this organism entered a new phase. Subsequently new tools were developed to speed up this process. Gene disruption is one of the most powerful techniques available for the study of gene function, and a breakthrough was achieved when it became apparent that only very short sequences of yeast DNA on either side of a marker gene are needed for efficient integration into the yeast genome by homologous recombination. This permits the use of gene disruption cassettes created by PCR (2,3). Thus, for the first time, large-scale, systematic gene disruption projects became feasible (4,5). The next important step was the introduction of the completely heterologous kanr marker, which confers resistance to the antibiotic G418 (6). This dominant marker dispenses with the need for yeast strains that are auxotrophic for the markers normally used for gene disruption. Moreover, because the kanr gene (which is driven by the Ashbya gossypiiTEF2 promoter and terminated by the A.gossypiiTEF2 terminator) shows no homology to the yeast genome, recombination between the yeast genome and internal parts of the kanMX disruption cassette (which would result in misintegration) is minimal, thus maximising the frequency of correct integration. Consequently, the kanMX gene disruption cassette was chosen for large-scale gene disruption projects, as well as for use in the genome-wide gene knockout project (5,7,8). In functional studies in S.cerevisiae it is often necessary to delete more than one gene. This is partly due to the fact that substantial portions of the yeast genome are duplicated (9,10). Moreover, as many cellular functions are maintained by more than one isoenzyme, it is often necessary to create multiple gene disruptions to uncover gene function (11–13). One prominent example is the hexose transporter family: in this case, concurrent knockout of at least 20 transporter genes was required to block uptake of hexose completely (14). Marker rescue and reuse offers a convenient and efficient way to introduce multiple gene disruptions. Removal of a previously integrated disruption cassette is most easily achieved if the respective cassette is flanked by directly repeated sequences 400–500 bp in length. Mitotic recombination between the repeated sequences results in excision of the marker and leaves a single repeat in the genome (15). This principle has been used with a variety of marker genes (16–20). The main problems encountered with this approach are: (i) the low rate of mitotic recombination, which makes it necessary to screen for rare instances of marker loss (except in cases where counter-selection is possible, e.g. URA3; 21); (ii) the fact that each marker rescue event leaves an additional repeat sequence in the genome which serves as an unwanted recombination target for integration of the gene disruption cassette in the next round of gene deletion. Recently we designed a new and efficient gene disruption cassette for repeated use by combining the advantages of the kanMX gene disruption cassette with those of the Cre/loxP recombination system of bacteriophage P1 (22). The kanMX cassette is flanked by two direct repeats of the 34 bp loxP sequence, yielding the loxP–kanMX–loxP cassette, which can be converted into a highly efficient gene disruption cassette by the addition of short DNA sequences homologous to the genomic locus of interest using PCR (Fig. 1). Once correctly integrated into the genome the kanMX marker can be efficiently rescued by transformation with a plasmid carrying the gene for Cre recombinase under control of the GAL1 promoter and induction of Cre expression by shifting the cells to galactose-containing medium. Cre-induced recombination results in loss of the kanMX cassette, leaving behind a single loxP site at the original integration site (Fig. 1) (22). Since its introduction the loxP–kanMX–loxP cassette has been widely used (see for example 23–26). Many gene families have now been analysed in this way: for example, the above mentioned hexose transporter family (14) and a series of gene families of unknown function investigated within the framework of the EUROFAN project (27; J.H.Hegemann and U.Gueldener, in preparation). Moreover, a modified loxP–kanMX–loxP cassette has been used to control gene expression in a novel way. For this purpose the loxP–kanMX–loxP cassette is embedded in the ACT1 intron and, after genomic integration of this ‘maxi-intron’ into the 5′-terminal region of the gene of interest, expression of this gene is completely inhibited. Gene expression is restored only after Cre-mediated site-specific recombination has deleted the kanMX unit and thus created a small spliceable intron carrying a single loxP site (28). In a different application the loxP–kanMX–loxP marker cassette was used to repeatedly add an epitope to different gene products at their C-termini (29). Finally the loxP–kanMX–loxP marker has been incorporated into an integration cassette that can be used to fuse GFP to any protein of interest (30; U.Gueldener and J.H.Hegemann, in preparation). Meanwhile the loxP–kanMX–loxP/Cre system has also been adapted successfully for use in the yeast Kluyveromyces lactis (31). One limitation of the Cre/loxP system so far has been its restriction to the kanr marker gene. This has meant that the system could not be used in the background of the kanMX deletion strains created in the genome-wide knockout project (8). Furthermore, it would be interesting in certain cases to follow a deleted gene or genes in genetic crosses (by following the disruption cassette markers) before the markers are excised by inducing Cre expression. In this paper we describe the construction of additional loxP–marker gene–loxP cassettes for repeated gene disruption. Most importantly we introduce a new dominant drug resistance cassette carrying the bler gene from the bacterial transposon Tn5, which confers resistance to the antibiotic phleomycin (Phleo). Finally, construction of additional Cre expression plasmids carrying HIS3, TRP1 or the dominant bler marker gene allows a much wider application of this powerful marker rescue system and significantly increases the number of new markers of the Cre/loxP system (22,27,31), thus further enhancing the utility of the Cre/loxP system in functional genome research. MATERIALS AND METHODS Strains and media Escherichia coli strain XL1-Blue was obtained from Stratagene (Heidelberg, Germany). The yeast strain CEN.PK2-1C was used for in vivo recombination to generate plasmid pSH65 and for gene disruption experiments. All culture media were prepared as described previously (32). For selection of G418 resistance after yeast transformations, cells were plated onto YPD plates containing 200 µg/ml G418 sulphate (Gibco BRL, Germany; batch 11811-031, activity may be batch-dependent) (dissolve in water and add to autoclaved YPD cooled to 60°C, to a final active concentration of 200 µg/ml) after an initial period of growth (2 h) in non-selective medium. Selection for phleomycin resistance was performed on YPD plates containing 7.5 µg/ml phleomycin (PHLEL0100; Cayla, France) (dissolve in water and add to autoclaved YPD cooled to 60°C). Systematic testing revealed that in this case preincubation in non-selective medium did not enhance transformation efficiency. For selection for the kanr gene in E.coli, bacteria were plated on YT plates containing 50 µg/ml kanamycin sulphate (M6750; Sigma, Germany) (6). Pfu polymerase (M7741) was obtained from Promega (Germany); Taq polymerase was prepared as described (33). Plasmid construction Positioning of new heterologous marker genes between twoloxPsites. Plasmid pUG6 carrying the loxP–kanMX–loxP module has been described previously (22). For construction of plasmid pUG27 (carrying loxP–his5+–loxP) the BglII–SacI DNA fragment from plasmid pFA6a-HIS3MX6 (18), carrying the Schizosaccharomyces pombehis5+ ORF (which complements the S.cerevisiaehis3 mutation) plus the A.gossypiiTEF2 promoter and terminator, was cloned into BglII + SacI-cleaved plasmid pUG6, replacing the kanMX unit between the two loxP sites. For construction of plasmid pUG66 (carrying loxP–bler–loxP) the vector pUT332, containing the bler gene from transposon Tn5 (34), was used as the template in a PCR with the oligonucleotide primers 1104 (5′-GCCGTAAGCCATGGCCGACCAAGCGACGCCCAAC-3′) and 1106 (5′-GGCGCCGGAGTACTGATCATGAGATGCCTGCAAGC-3′) (restriction sites underlined) to amplify the bler open reading frame. The resulting PCR product was cut with NcoI and ScaI and ligated with NcoI + ScaI-cleaved pUG6, thus replacing the kanr coding sequence. For construction of plasmid pUG72 (carrying loxP–URA3–loxP; originally named pJJH726) the K.lactisURA3 gene including the promoter and terminator was amplified from genomic DNA of the K.lactis type strain CBS2359 with oligonucleotides KlURA3-SacI (5′-GGCGAGCTCGTTTTATTTAGGTTCTATCGAGG-3′) and KlURA3-BamHI (5′-CCGCGGATCCCAATACAACAGATCACGTG-3′). The resulting PCR product of ∼1.4 kb was cleaved with SacI and BamHI and ligated to BglII + SacI-cleaved pUG6, thus replacing the complete kanMX unit. For construction of plasmid pUG73 (carrying loxP–LEU2–loxP; originally named pJJH727) the K.lactisLEU2 gene plus promoter and terminator sequences was amplified by PCR using K.lactis genomic DNA and the oligonucleotides KlLEU2-SacI (5′-GGCGAGCTCGCTGTGAAGATCCCAGCAAAGG-3′) and KlLEU2-BamHI (5′-GGCGGGATCCGCAGGCTAACCGGAACCTG-3′). The PCR product of ∼2.2 kb was digested with SacI and BamHI and ligated into BglII + SacI-cut pUG6. Construction of new Cre expression plasmids. Plasmid pSH47 (GAL1-cre, URA3) has been described previously (22). For construction of plasmid pSH62 (GAL1-cre, HIS3) a 1.4 kb EcoRI–XhoI DNA fragment of pSH47 carrying the cre open reading frame was ligated with EcoRI + XhoI-cleaved p413GAL1 (35). For construction of plasmid pSH63 (GAL1-cre, TRP1) the 1.4 kb EcoRI–XhoI DNA fragment of pSH47 carrying the cre gene was ligated to EcoRI + XhoI-cut p414GAL1 (35). For construction of plasmid pSH65 (GAL1-cre, ble) the vector pUT332, containing the bler gene from transposon Tn5 plus the S.cerevisiaeTEF1 promoter (34), was used as template in a PCR with the oligonucleotide primers 1095 (5′-AGAAGGTTTTGGGACGCTCGAAGGCTTTAATTTGCGGCCGCGTTGTAAAACGACGGCCAG-3′) and 1096 (5′-AGTGAGCGCGCGTAATACGACTCACTATAGGGCGAATTGGGCGTACACGCGTCTGTACAG-3′). At their 3′-ends the oligonucleotides (underlined) are homologous to plasmid pUT322, while the 5′-ends of the oligonucleotides are homologous to the sequences to the left and right of the KpnI restriction site in pSH47. The ∼1.0 kb PCR product carrying the bler gene plus the S.cerevisiaeTEF1 promoter was transformed into yeast strain CEN.PK2 together with the KpnI-cleaved plasmid pSH47, selecting for uracil prototrophy after homologous recombination of both DNA molecules in yeast. Genomic DNA from individual yeast transformants was isolated (32), transformed into E.coli and plasmid DNA was prepared. The correct plasmid, called pSH64, was identified by restriction enzyme and sequence analysis. Subsequently pSH64 was cut with NcoI (site located within the URA3 coding sequence) and the ends were filled-in with Pfu polymerase and religated to inactivate the URA3 gene, yielding pSH65. The complete DNA sequences of all pUG and pSH plasmids have been deposited in GenBank and the corresponding accession numbers can be found in Figures 2 and 5. Plasmid requests The vectors pUG6 (kanr), pUG27 (his5+), pUG66 (bler), pUG72 (URA3) and pUG73 (LEU2) carrying the various loxP–marker gene–loxP gene disruption cassettes as well as the Cre expression plasmids pSH47 (yeast selection marker URA3), pSH62 (HIS3), pSH63 (TRP1) and pSH65 (bler) can be obtained individually (pUG6, accession no. P30114; pUG27, accession no. P30115; pUG66, accession no. P30116; pUG72, accession no. P30117; pUG73, accession no. P30118; pSH47, accession no. P30119; pSH62, accession no. P30120; pSH63, accession no. P30121; pSH65, accession no. P30122) or as a package (DEL-MARKER-SET) from Euroscarf (Frankfurt, Germany; http://www.uni-frankfurt.de/fb15/mikro/euroscarf/). The package includes the heterologous gene disruption cassettes from the McCusker group (19,20). Companies should contact Johannes H. Hegemann. PCR-mediated generation of loxP–marker gene–loxP gene disruption cassettes and their integration into the yeast genome All loxP–marker gene–loxP disruption cassettes presented in this work can be generated by PCR using oligonucleotides carrying the same 19 and 22 3′ nucleotides, while the 40 5′ nucleotides must be homologous to sequences to the left or right of the gene to be deleted (see Fig. 2 and Table 1). As a test case for the new disruption cassettes the ADE2 and YOR387c genes were deleted using the oligonucleotides listed in Table 1. For each disruption cassette one preparative PCR was performed as described previously (22). The PCR conditions were: 95°C for 5 min (hot start); 25 cycles of 94°C for 40 s, 58°C for 1 min and 68°C for 2 min; a final step at 68°C for 15 min. The PCR product was purified and the cassettes used for yeast transformation as described before (22). RESULTS AND DISCUSSION Construction of new loxP-flanked gene disruption cassettes The recently introduced loxP–kanMX–loxP gene disruption cassette combines the advantages of the heterologous, dominant kanr marker with the capacity for efficient gene disruption by Cre-mediated recombination between the two loxP sites, resulting in efficient marker rescue (22) (Fig. 1). In order to expand the Cre/loxP technology and maximise the utility of this system, we have constructed additional loxP–marker gene–loxP cassettes. The four newly created loxP-flanked marker cassettes, like the original loxP–kanMX–loxP cassette, completely lack homology to the genome of the budding yeast, which maximises the probability of recovering the desired homologous integration event (Fig. 2). Construction of the new gene disruption cassettes started from the widely used loxP–kanMX–loxP cassette cloned in plasmid pUG6 (22). For construction of the loxP–his5+–loxP cassette the kanMX unit located between the two loxP sites in plasmid pUG6 was replaced by the S.pombehis5+ unit obtained from plasmid pFA6a-HIS3MX6 (18), yielding plasmid pUG27. The S.pombehis5+ gene complements the S.cerevisiaehis3 mutation. The overall degree of identity between the S.pombehis5+ and S.cerevisiaeHIS3 DNA sequences is 59%, with several blocks exhibiting higher homology (70–76% identity). This, however, is too low to allow efficient homologous integration of the S.pombe gene at the HIS3 locus (18). For construction of the heterologous loxP–URA3–loxP and loxP–LEU2–loxP cassettes, both genes plus their promoter and terminator sequences were isolated by PCR from K.lactis genomic DNA and cloned between the two loxP sequences in plasmid pUG6, replacing the kanMX unit and resulting in the plasmids pUG72 and pUG73. The two K.lactis genes complement the corresponding ura3 and leu2 mutations in S.cerevisiae (36,37). The K.lactis genes are 73 (URA3) and 77% (LEU2) identical to their S.cerevisiae homologues. This is too low to permit efficient recombination between the corresponding K.lactis and S.cerevisiae genes, in particular as the longest stretch of DNA sequence identity between corresponding genes is 17 bp in the case of URA3 and 18 bp for LEU2. The loxP–URA3–loxP cassette should be particularly useful, because (i) most S.cerevisiae laboratory strains are ura3– and (ii) 5-fluoroorotic acid-containing media can be readily used for counter-selection (21). Finally, the bler gene from transposon Tn5 was used to create the new heterologous dominant drug resistance marker cassette loxP–bler–loxP. The bler gene renders prokaryotes and lower eukaryotes resistant to phleomycin, which belongs to the metallo-glycopeptide group of antibiotics of the bleomycin family (38). Cells expressing the bler gene become resistant to phleomycin as a result of tight binding of the antibiotic by the Ble protein. The Ble protein appears not to be toxic to S.cerevisiae. Phleomycin is produced in fermentation cultures by Streptomyces verticillus (39). It has previously been shown that expression of the bler gene from an episomal or an integrative plasmid allows direct selection of phleomycin-resistant S.cerevisiae transformants (34,38,40). The bler gene was isolated by PCR from plasmid pUT332 (34) and placed between the A.gossypiiTEF promoter and terminator in plasmid pUG6, yielding plasmid pUG66 (Fig. 2). Gene disruption properties of the new loxP–marker gene–loxP cassettes An important property of the new cloned gene disruption cassettes is that, like the original loxP–kanMX–loxP cassette (22), each of the marker genes plus regulatory elements is flanked by two loxP sequences embedded in the same vector backbone (see Fig. 2 and Materials and Methods). This allows the use of the same primer sequences to amplify all of the gene disruption cassettes. To test the fidelity of the new cassettes for targeted gene disruption all cassettes were used to delete two genes: ADE2 and YOR387c. The 2 × 5 gene disruption cassettes were amplified in parallel using primers 1134 and 1135 (for ADE2) and 1017 and 1018 (for YOR387c) (Table 1). The 5′-ends of the primers contain 45 nt stretches that are homologous to sequences upstream of the ATG start codon and downstream of the stop codon, respectively, of the target genes, followed by a 19 nt segment homologous to sequences to the left of the loxP site and a 22 nt motif that anneals to the right of the loxP site of the disruption cassettes (Figs 1 and 2). Comparable amounts of the various disruption cassettes were generated by PCR and purified (Fig. 3). Approximately 2 µg of the gene disruption cassettes were transformed into strain CEN.PK2-1C and transformants were selected on the corresponding SD dropout plates, on YPD + 7.5 µg/ml phleomycin (for the loxP–bler–loxP cassette) or on YPD + 200 µg/ml G418 (for the loxP–kanMX–loxP cassette) plates, as described before (22). Transformation of the five different disruption cassettes in all cases yielded comparable numbers of transformants. For selection of phleomycin-resistant colonies we first determined the usable range of antibiotic concentrations and found that 7.5 µg/ml phleomycin (in YPD) was optimal for selection of loxP–bler–loxP transformants. At this concentration no spontaneous phleomycin-resistant colonies were found. In contrast to what has been described for transformation with a bler gene-carrying multicopy plasmid (40), we could not improve the transformation efficiency of the loxP–bler–loxP cassette further by incubating the transformation mix in liquid YPD medium for several hours (up to 10 h was tested) prior to plating on YPD plates containing phleomycin. However, we did not try to enhance the transformation efficiency of the loxP–bler–loxP cassette further by adding glycerol to the transformation plates (38). To confirm correct integration of the various disruption cassettes into the ADE2 and YOR387c loci, 24 transformants from each plate were subjected to colony PCR using combinations of the corresponding target gene-specific (primers A–D) and disruption cassette-specific primers (primers C-B and C-C) (Fig. 1 and Table 1). A typical example of the results of the PCR analysis of putative yor387c deletants using target gene-specific primer A and one of the three cassette-specific primers C-B is shown in Figure 4. The PCR products obtained from the various deletion strains are between 350 and 600 bp in length and thus exhibit the calculated cassette-specific sizes predicted for correct integration. The frequency of correctly integrated cassettes at the YOR387c locus varied between 46% for the loxP–kanMX–loxP cassette and 100% for the loxP–his5+–loxP and loxP–LEU2–loxP cassettes. Analysis of the ade2 transformants revealed a similar level of correctly integrated cassettes: between 52% (loxP–kanMX–loxP) and 92% (loxP–URA3–loxP) of the transformants had undergone the correct homologous recombination event (Fig. 4). As expected, all ade2 deletion strains characterised by PCR also produced red colonies on indicator plates due to accumulation of a biosynthetic intermediate product, thus confirming the PCR results (data not shown). In summary, PCR analysis of putative yor387c and ade2 deletants revealed that all five cassettes disrupted their target genes with similar efficiency: at least 50% of the transformants had undergone the predicted homologous recombination event. Similar efficiencies for this type of gene disruption have been observed by others, mainly due to the fact that the gene disruption cassettes used in these cases consist of completely heterologous DNA sequences (6,19,22). Our results show that the two heterologous K.lactis genes URA3 and LEU2 exhibit excellent selection marker qualities for this type of gene deletion. In fact, the URA3 gene from this yeast has previously been used for gene disruption experiments by others (27,41). Likewise, the his5+ gene from S.pombe proved to be an efficient marker for gene disruption, as has been reported by others for a non-loxP version of the his5+ gene (18,42). The results of the gene knockout experiment show that the bler gene is an efficient heterologous dominant resistance marker: 63 and 96% of the transformants proved to have correctly integrated the loxP–bler–loxP gene disruption cassette at the ADE2 and YOR387c loci, respectively. The transformation efficiency of this cassette is comparable to that of the second dominant resistance marker, loxP–kanMX–loxP, and to that of the other heterologous auxotrophic markers used here (data not shown). Moreover, ade2Δ strains carrying the bler marker showed no obvious growth disadvantage when compared with ade2Δ strains containing one of the other marker genes, suggesting that these markers are probably phenotypically neutral. However, this needs to be tested more thoroughly in the future. To exclude the possibility that high level expression of the phleomycin resistance protein has adverse effects on cellular functions in S.cerevisiae, the marker can be easily removed by activating the Cre-mediated recombination process to delete the bler gene. Finally, it is important to note that no cross-resistance with other currently used dominant resistance markers has been reported. Therefore, phleomycin can be used to select cells that are already resistant to other selective agents, for example G418. This makes the dominant loxP–bler–loxP cassette a very useful tool when working with the kanr-based KO strain collection produced by the Transatlantic Yeast Consortium during the EU project EUROFAN 2. Our laboratory routinely uses both dominant markers to produce double deletion strains. New Cre expression plasmids for marker rescue Expression of Cre recombinase in yeast strains carrying one (or more) of the loxP–marker gene–loxP cassettes results in efficient recombination between the loxP sites, removing the marker gene and leaving behind a single loxP site at the chromosomal locus (22). To increase the flexibility of this system, we have exchanged the URA3 selection marker in the original cre expression vector pSH47 for HIS3 (resulting in plasmid pSH62) and TRP1 (pSH63), as well as for the dominant resistant marker bler (pSH65) (Fig. 5). The cre expression cassette consisting of the inducible GAL1 promoter, the cre gene and the CYC1 terminator is not affected by these modifications. The new vectors were tested for efficiency of marker rescue and were found to induce the same rates of marker loss as described for the original pSH47 vector (22; data not shown). In the meantime these new cre expression vectors have also been successfully used by others (see for example 28). In our routine work we no longer induce Cre expression by shifting loxP–marker gene–loxP marked cells transformed with a cre plasmid to galactose medium. Instead, we grow these cells overnight in liquid YPD medium (with glucose as carbon source) and then plate ∼200 cells on YPD plates. Replica plating onto YPD and YPD plus the drug (or onto the corresponding selective medium in the case of the auxotrophic markers) results in the identification of cells which have lost the marker gene from the disruption cassette (∼5% of the total). This is probably due to residual activity of the GAL1 promoter in the presence of glucose. Several loxP-flanked marker genes can be removed from the genome simultaneously (data not shown). However, care must be taken not to induce Cre expression too strongly (e.g. by using multicopy plasmids or inducing Cre expression for prolonged times), as this will result in unintended recombination events between various loxP sites (27). Swapping of cassette markers An additional advantage of the loxP-flanked kanMX, bler and his5+ marker genes is their use for marker cassette exchange. The presence of completely identical DNA sequences of ∼300 bp to the left and right of the marker gene in each cassette (see Fig. 2) allows for easy replacement of each of the marker genes by any of the others. For this purpose only two oligonucleotides have to be designed which allow amplification of the marker gene of interest plus the promoter, terminator and loxP sequences. Transformation of this cassette into a yeast strain carrying one of the other two cassettes integrated into the genome replaces the marker gene within the integrated cassette. Successful cassette swapping can be verified by PCR analysis using target gene-specific (A and D) and new cassette-specific primers. To exclude unintended targeting events, swapping experiments should be restricted to single deletion strains that do not carry plasmid pSH65 (carrying S.cerevisiaeTEF1 sequences). Conclusion We have constructed four new gene disruption cassettes with the marker genes bler, his5+, URA3 and LEU2, consisting of completely heterologous DNA flanked by loxP sequences. In addition, the Cre expression plasmid has been equipped with three other selection markers (HIS3, TRP1 and bler). The set of five different gene disruption cassettes and four different Cre expression vectors presented here will significantly extend the tool kit for functional analysis of the S.cerevisiae genome. In fact, some of the new vectors have already been used successfully in a variety of experiments (see for example 28,43; J.Heinisch, unpublished results). The availability of the two dominant marker genes kanr and bler and a Cre expression vector carrying bler as the yeast selection marker facilitates the creation of multiple gene knockouts not only in commonly used laboratory strains, but also in prototrophic industrial or wild-type strains of S.cerevisiae. ACKNOWLEDGEMENTS We thank Ute Gengenbacher and Sabine Klein for technical assistance. Dr Paul Hardy is thanked for critical reading of the manuscript. We are grateful to Dr Yde Steensma for plasmid pUT332. This work was supported by a contract awarded to J.H.H. in the context of the EUROFAN project of the EC (BIO4-CT95-0080) and a grant from the Deutsche Forschungsgemeinschaft (He1880/4-1) to J.H. * To whom correspondence should be addressed. Tel: +49 211 81 13733; Fax: +49 211 81 13567; Email: [email protected] Present addresses:U. Gueldener, MIPS, GSF, Ingolstaedter Landstrasse 1, 85764 Neuherberg, GermanyJ. Heinisch, Universität Hohenheim, Institut für Lebensmitteltechnologie (150), Fachgebiet Gärungstechnologie, Garbenstrasse 25, 70599 Stuttgart, Germany +AF298780, AF298782, AF298785, AF298788–AF298790, AF298792–AF298794 View largeDownload slide Figure 1. The loxP/Cre gene disruption and marker rescue procedure. The gene disruption cassette consists of a selection marker gene (marker), usually conferring drug resistance or prototrophy, flanked by two 34 bp loxP sequences as direct repeats located adjacent to 45 bp of sequence flanking the chromosomal target sequence (ORF) to be deleted. The disruption cassette is produced by PCR using oligonucleotides comprising 19 or 22 3′ nucleotides complementary to sequences in the template (marker plasmid) flanking the disruption cassette and 45 5′ nucleotides that anneal to sites upstream or downstream of the genomic target sequence to be deleted. After transformation of the linear disruption cassette into yeast cells, selected transformants are checked by PCR for correct integration of the cassette and concurrent deletion of the chromosomal target sequence. The verification PCRs are done using combinations of primers complementary to sequences within the cassette (C-B, cassette B primer; C-C, cassette C primer) and to sequences within or flanking the target sequence (A, B, C and D). In a diploid yeast cell (shown here) in which one allele of the target sequence has been replaced by a disruption cassette, all PCRs indicated will produce products of the expected size. Finally, expression of the Cre recombinase results in removal of the marker gene, leaving behind a single loxP site at the chromosomal locus. View largeDownload slide Figure 1. The loxP/Cre gene disruption and marker rescue procedure. The gene disruption cassette consists of a selection marker gene (marker), usually conferring drug resistance or prototrophy, flanked by two 34 bp loxP sequences as direct repeats located adjacent to 45 bp of sequence flanking the chromosomal target sequence (ORF) to be deleted. The disruption cassette is produced by PCR using oligonucleotides comprising 19 or 22 3′ nucleotides complementary to sequences in the template (marker plasmid) flanking the disruption cassette and 45 5′ nucleotides that anneal to sites upstream or downstream of the genomic target sequence to be deleted. After transformation of the linear disruption cassette into yeast cells, selected transformants are checked by PCR for correct integration of the cassette and concurrent deletion of the chromosomal target sequence. The verification PCRs are done using combinations of primers complementary to sequences within the cassette (C-B, cassette B primer; C-C, cassette C primer) and to sequences within or flanking the target sequence (A, B, C and D). In a diploid yeast cell (shown here) in which one allele of the target sequence has been replaced by a disruption cassette, all PCRs indicated will produce products of the expected size. Finally, expression of the Cre recombinase results in removal of the marker gene, leaving behind a single loxP site at the chromosomal locus. View largeDownload slide Figure 2. The series of loxP–marker gene–loxP gene disruption cassettes. The marker plasmids pUG6, pUG27, pUG66, pUG72 and pUG73 serve as templates for PCR to generate the five different gene disruption cassettes. All marker genes (selectable genes) are derived from organisms other than S.cerevisiae and are expressed from the A.gossypiiTEF2 promoter and terminated by the TEF2 terminator (6), except for the two K.lactis genes, URA3 and LEU2, which retain their own regulatory sequences. kanr and bler are dominant resistance marker genes conferring resistance to G418 and phleomycin (selectable phenotype), respectively. Each marker gene including promoter and terminator is flanked by loxP sites. Since the vector backbone is the same in all cases, each of the five disruption cassettes can be amplified by PCR using the same two primers. The complete vector sequences have been deposited at GenBank under the accession numbers listed in the figure (Sequence db acc. #). View largeDownload slide Figure 2. The series of loxP–marker gene–loxP gene disruption cassettes. The marker plasmids pUG6, pUG27, pUG66, pUG72 and pUG73 serve as templates for PCR to generate the five different gene disruption cassettes. All marker genes (selectable genes) are derived from organisms other than S.cerevisiae and are expressed from the A.gossypiiTEF2 promoter and terminated by the TEF2 terminator (6), except for the two K.lactis genes, URA3 and LEU2, which retain their own regulatory sequences. kanr and bler are dominant resistance marker genes conferring resistance to G418 and phleomycin (selectable phenotype), respectively. Each marker gene including promoter and terminator is flanked by loxP sites. Since the vector backbone is the same in all cases, each of the five disruption cassettes can be amplified by PCR using the same two primers. The complete vector sequences have been deposited at GenBank under the accession numbers listed in the figure (Sequence db acc. #). View largeDownload slide Figure 3. For deletion of the ADE2 gene, five different gene disruption cassettes were produced in parallel PCRs using the primers 1134 and 1135 (see Table 1). The sizes of the cassettes vary between 1300 and 2500 bp (2% of each PCR product was loaded). View largeDownload slide Figure 3. For deletion of the ADE2 gene, five different gene disruption cassettes were produced in parallel PCRs using the primers 1134 and 1135 (see Table 1). The sizes of the cassettes vary between 1300 and 2500 bp (2% of each PCR product was loaded). View largeDownload slide Figure 4. PCR analysis to confirm correct integration of the gene disruption cassettes at the YOR387c and ADE2 loci (see Fig. 1). PCR was carried out on transformants obtained in the YOR387c disruption experiment, using target gene-specific primer A (primer 1025) and the disruption cassette-specific primer C-B (for kanr, his5+ and bler the kan-B primer 363; for URA3 the Ura-B primer 1115; for LEU2 the Leu2-B primer 1136). The products obtained from two yor387cΔ mutants (Δ1 and Δ2) disrupted with each of the various disruption cassettes and from wild-type (wt) cells were then subjected to agarose gel electrophoresis. For both genes 24 transformants obtained with each of the gene disruption cassettes were analysed in similar experiments. The percentages of correctly integrated cassettes at YOR387c and ADE2 are indicated below the gel. DNA fragment size markers: M1, EcoRI + HindIII-cleaved λ DNA; M2, Bsp143I-cleaved pUC19 (the sizes of selected marker DNA fragments are shown on the right). View largeDownload slide Figure 4. PCR analysis to confirm correct integration of the gene disruption cassettes at the YOR387c and ADE2 loci (see Fig. 1). PCR was carried out on transformants obtained in the YOR387c disruption experiment, using target gene-specific primer A (primer 1025) and the disruption cassette-specific primer C-B (for kanr, his5+ and bler the kan-B primer 363; for URA3 the Ura-B primer 1115; for LEU2 the Leu2-B primer 1136). The products obtained from two yor387cΔ mutants (Δ1 and Δ2) disrupted with each of the various disruption cassettes and from wild-type (wt) cells were then subjected to agarose gel electrophoresis. For both genes 24 transformants obtained with each of the gene disruption cassettes were analysed in similar experiments. The percentages of correctly integrated cassettes at YOR387c and ADE2 are indicated below the gel. DNA fragment size markers: M1, EcoRI + HindIII-cleaved λ DNA; M2, Bsp143I-cleaved pUC19 (the sizes of selected marker DNA fragments are shown on the right). View largeDownload slide Figure 5. Cre expression plasmids. All plasmids carry the CEN6/ARSH4 module to ensure stability in yeast and a Cre expression cassette consisting of the cre open reading frame flanked by the galactose-inducible GAL1 promoter and the CYC1 terminator from S.cerevisiae. The vectors differ in the type of selection marker used. The URA3, HIS3 and TRP1 marker genes are from S.cerevisiae and retain their own regulatory sequences, while the bler gene from transposon Tn5 is expressed from the S.cerevisiaeTEF1 promoter. View largeDownload slide Figure 5. Cre expression plasmids. All plasmids carry the CEN6/ARSH4 module to ensure stability in yeast and a Cre expression cassette consisting of the cre open reading frame flanked by the galactose-inducible GAL1 promoter and the CYC1 terminator from S.cerevisiae. The vectors differ in the type of selection marker used. 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El-Maarri, Osman;Herbiniaux, Ursula;Walter, Jörn;Oldenburg, Johannes

doi: 10.1093/nar/30.6.e25pmid: 11884644

Abstract The precise mapping and quantification of DNA methylation as an epigenetic parameter during development and in diseased tissues is of great importance for functional genomics. Here we describe a rapid, quantitative method to assess methylation levels at specific CpG sites using PCR products of bisulfite-treated genomic DNA. Using single nucleotide primer extension (SNuPE) assays in combination with ion pair reverse phase high performance liquid chromatography (IP RP HPLC) separation techniques, methylated and unmethylated CpGs can be discriminated and quantified based on the different masses and hydrophobicities of the extended primer products. The assay is linear, highly reproducible and several sites can be measured simultaneously in one reaction. It can be semi-automated and eliminates the need for cloning and sequencing of individual bisulfite PCR products. Received November 15, 2001; Revised and Accepted January 29, 2002. INTRODUCTION Epigenetic control of gene expression plays a crucial role in functional genomics. The precise measurement of epigenetic changes is of particular interest in the fields of developmental biology, cancer, multifactorial diseases and aging. Many methods have been developed for the detection of methylation, among which the bisulfite-based sequencing technique is the most precise and sensitive, providing sequence-independent information (1,2). A number of successful modifications of the original protocols have been published, mostly improving technical handling and sensitivity of the method. However, little progress has been made to circumvent the laborious steps of cloning and sequencing (3,4). Although a bisulfite-based single nucleotide primer extension (SNuPE) assay has been described for direct quantification, this protocol still requires radioactive quantification following gel separation and hence cannot be applied for high throughput performance (5). Here we present a simple SNuPE assay in combination with an ion pair reverse phase high performance liquid chromatography (IP RP HPLC) separation method to rapidly quantify the methylation status at several CpG sites simultaneously. The basis for the method is a combination of bisulfite PCR modification, primer extension and IP RP HPLC separation of the extended primer products. After bisulfite treatment of the DNA (5) and PCR amplification of the region to be analyzed, the PCR products are purified to remove non-extended oligos and dNTPs. Following the purification step, specific primers just 5′ to a CpG site(s) are hybridized to the denatured single-stranded PCR product. The annealed primers are then extended using Sequenase in the presence of both ddCTP and ddTTP. The incorporation of ddTTP indicates that the original chromosomal DNA is unmethylated and conversely the incorporation of ddCTP indicates that the CpG is methylated (Fig. 1). In order to quantitate the incorporation of either ddTTP or ddCTP the extension products are directly (without any further purification) loaded on a dHPLC column (WAVE DNA Fragment Analysis System; Transgenomics). The amounts of ddTTP and ddCTP extended products are quantified by integrating the areas of the peaks and calculation of their percentage ratios using the WaveMaker program v.4.1 (Transgenomics) (see also Materials and Methods). MATERIALS AND METHODS Bisulfite treatment Bisulfite treatment of DNA was performed by a bead-based method as described by Engemann et al. (6). The primers used for amplification of bisulfite-treated lymphocyte DNA from Factor VIII exon 14 containing eight CpG sites at codons 1602, 1620, 1645, 1648, 1652, 1689 and 1696, were as described by El-Maarri et al. (7), while the primers used to amplify the 5′‐region of the SNRPN gene were as described by El-Maarri et al. (2). Primer extension reaction Primers used for the primer extension reaction were as follows: SN-F8-1645, 5′-aat tta tta gtt ttg aaa-3′; SN-F8-1689, 5′-gat gaa aat tag agt ttt-3′; SN-F8-1696, 5′-ttt ttt agt ttt taa aag aaa ata-3′; SN-SNRPN-2-3t, 5′-ttt ggg att ttt gta ttg-3′; SN‐SNRPN-21, 5′-taa ggt tag ttg tgt-3′; SN-SNRPN-23, 5′-ga tag ttt ggg gag-3′. The underlined t residues do not correspond to the DNA sequence and were added to increase the masses and hydrophobicities and thus the retention times of these oligos. The primers used in the SNuPE reactions are 5′ to CpG sites in codons 1645, 1689 and 1696 of the Factor VIII gene and CpG sites 2, 21 and 23 in the 5′-region of the SNRPN gene, as described by El-Maarri et al. (2). The reaction was carried out as described by Hoogendoorn et al. (8). Briefly the reaction was carried out in 20 µl total volume containing ∼50 ng purified PCR product, 50 µM each ddCTP and ddTTP (Amersham), 12.5 pmol each primer and 3 U ThermoSequenase (Amersham). The thermal cycling was as follows: initial denaturation step of 3 min at 95°C, followed by 50 cycles of 30 s at 42°C and 2 min at 60°C. dHPLC analysis The HPLC analysis was performed on a WAVE DNA Fragment Analysis System from Transgenomics. An aliquot of 10 µl of the SNuPE reaction was loaded. The oven temperature was set to 50°C and elution was with a gradient of acetonitrile made by mixing buffers A and B, consisting of 0.1 M triethyammonium acetate (TEAA) buffer and 0.1 M TEAA buffer with 25% acetonitrile, respectively. Each run was made up of a gradient of 30–41% buffer B for the Factor VIII gene and 24–43% buffer B for the SNRPN gene over 10 min, followed by re-equilibration of the column by injection of 30% buffer B for 1 min. The eluted DNA was detected with a UV detector at 260 nm. Calculation of methylation extent The degree of methylation at a given CpG site was determined according to the term %Meth = (AC/AC + AT) × 100, where AC and AT are the surface areas under the peaks corresponding to the ddCTP-extended primer and the ddTTP-extended primer, respectively, as calculated by Wave Maker v.4.1 (Transgenomics). RESULTS At 50°C, using TEAA as the ion-pairing reagent, separation of short extended oligos by dHPLC is based on differences in both their length (mass) and hydrophobicity. As a consequence the incorporation, during the SNuPE reaction, of the more hydrophilic ddTTP increases the retention time compared to oligos extended by ddCTP (Fig. 1). Therefore, the order of appearance of the oligos is unextended oligo followed by the ddCTP- and ddTTP-extended oligos, respectively. Separation of the above three oligos is good enough to give three distinguishable peaks that can be accurately measured. To test the accuracy and linearity of the reaction, PCR templates of the Factor VIII exon 14 region were analyzed as serial mixtures of methylated and unmethylated products, with a 10% increment in methylation (Fig. 2). Concordant with the expected 10% increment, the determined values exhibited a striking linearity for all three CpG sites studied. The slight non-linearity determined for one of the primer products may be due to increased non-specific annealing of its T-rich 3′-end to the rather A-rich sequence stretches of the bisulfite-treated PCR template. To avoid mispriming we recommend that the PCR fragment generated should be kept as short as possible and that the annealing and extension temperatures are optimized. As a second test for the accuracy of our approach we analyzed different PCR products derived from the 5′-region of SNRPN exon 1. Using the SNuPE-IP RP HPLC assay we found that the methylation levels at three selected CpG sites were 33, 40 and 42%, respectively (average of six different samples, data not shown; see Fig. 1B for an example). These values were concordant with previous results obtained using cloning and sequencing approaches, which showed that the majority of CpGs in this region are methylated on ∼40% of the chromosomes (2). Initially all three primers used for analysis of the CpG sites in Factor VIII exon 14 (1645, 1689 and 1696) were designed with an identical length of 18 bases. Based on their different base compositions (thymines) the corresponding retention times were as follows: SN-F8-1645, 50% thymines, 5.36 min; SN-F8-1689, 36% thymines, 1.55 min; SN-F8-1696, 33% thymines, 2.81 min. One problem encountered in a parallel multiplex analysis was that the products of primers SN-F8-1689 and SN-F8-1696 partially co-migrated and clear separation and quantification of the peaks was impossible (data not shown). To overcome this problem, primers of variable length and hydrophobicity were designed. The addition of six thymidines to the 5′-end of oligo SN-F8-1696 shifted its retention time from 2.81 to 8.16 min under the same assay conditions, resulting in clean separation of the extended products (Fig. 3). We therefore recommend this modification as a general strategy to facilitate multiplex column separation of the SNuPE products. DISCUSSION In recent years the bisulfite-based sequencing method has been by far the most widely used technique to analyze chromosomal methylation patterns. It provides sequence-independent information about the degree of methylation at a given CpG site and is sensitive enough to study methylation patterns in only a few cells. However, one major drawback of this method is that it requires laborious and time consuming cloning and sequencing steps. The described SNuPE-IP RP HPLC approach circumvents these laborious steps and provides a quick quantitative assay for a limited number of CpGs in the region of interest. The method will be extremely helpful to scrutinize methylation changes in larger numbers of samples at CpGs which are known to play a critical role. Since both the SNuPE-IP RP HPLC assay and the conventional cloning/sequencing method require complete bisulfite conversion and unbiased PCR amplification, a general problem for both methods may occur when using small amounts of starting material. Whereas the completeness of bisulfite conversion can be optimized by introducing several modifcations (6), a bias in the analysis may still arise when only a limited number of chromosomes are successfully amplified and hence the determined methylation levels do not represent the true methylation levels of all chromosomes. This can only be avoided by comparing results from independent bisulfite and PCR assays. Again, in this respect the SNuPE-IP RP HPLC approach would give a quick estimate of the reproductiveness of the different experiments. Moreover, this method could also be used to determine the parental origin of methylation, however, two restrictions apply. The first is, as for all traditional methylation analysis methods, a known polymorphism is required. The second is that it should be in the vicinity of a CpG site. When this is the case, two oligos should be designed that correspond to the two alleles of the polymorphism, with about 5–7 bases flanking a CpG site. If the site is differentially methylated then only one of the two oligos will be extended by ddCTP (for methylated) and the other by ddTTP (unmethylated). When performing the SNuPE-IP RP HPLC assay special attention should be given to primer design. Optimal primers should not be too short (between 15 and 18 bases) and should have a similar Tm of ∼40°C. Generally primers with degenerate positions (e.g. C or T) should be avoided. This causes difficulties in designing primers in CpG-dense regions, as the closeness of two successive CpGs may impede primer design. However, our data on the SNRPN region show that even in a CpG island, optimal primers can be selected and data comparable to cloning/sequencing analysis can be obtained. Furthermore, the content of T residues in the primer design should be carefully considered. Multiplexed primers should have different T residue contents, resulting in different hydrophobicities and column separation. If this cannot be done because of sequence constraints in the template we recommend artificially varying the hydrophobicity by adding stretches of T residues at the 3′‐ends of the primers. In conclusion, we have demonstrated that SNuPE-IP RP HPLC can be applied for the rapid analysis of methylation differences at specific CpG sites urgently needed in studies of carcinogenesis, imprinting diseases and aging. The assay is highly beneficial in comparing large numbers of samples in a relatively very short time. In combination with differential fluorescence labeling of the oligos, the assay will allow further multiplexing and facilitate the analysis of a substantial number of CpG sites in large series of samples in a short time at relatively low cost. ACKNOWLEDGEMENT Part of this work was supported by a BONFOR grant (O‐145.0004). * To whom correspondence should be addressed. Tel: +49 228 287 6737; Fax: +49 228 287 4320; Email: [email protected] View largeDownload slide Figure 1. Chromatograms of the multiplexed SNuPE reaction. (A) In the Factor VIII exon 14 region, representing a theoretical 50% methylated and 50% unmethylated. (B) In the SNRPN region. For each of the oligos used the unextended primer is indicated by a vertical arrow followed by the +C and +T extended primers representing the methylated (extended by ddCTP) and the unmethylated (extended by ddTTP) portions, respectively. All unlabeled peaks represent impurities from the initial unreacted primers. View largeDownload slide Figure 1. Chromatograms of the multiplexed SNuPE reaction. (A) In the Factor VIII exon 14 region, representing a theoretical 50% methylated and 50% unmethylated. (B) In the SNRPN region. For each of the oligos used the unextended primer is indicated by a vertical arrow followed by the +C and +T extended primers representing the methylated (extended by ddCTP) and the unmethylated (extended by ddTTP) portions, respectively. All unlabeled peaks represent impurities from the initial unreacted primers. View largeDownload slide Figure 2. The linearity curves for the three primers used in the SNuPE reaction (SN-F8-1645, SN-F8-1689 and SN-F8-1696-6t). The standard deviation of the data is shown for each measurement by a vertical bar. The theoretical equation of the linear response is shown above each graph. View largeDownload slide Figure 2. The linearity curves for the three primers used in the SNuPE reaction (SN-F8-1645, SN-F8-1689 and SN-F8-1696-6t). The standard deviation of the data is shown for each measurement by a vertical bar. The theoretical equation of the linear response is shown above each graph. View largeDownload slide Figure 3. The retention times of the SN-F8-1696 oligos: (a) unmodified; (b) with six A residues at the 5′-end; (c) with six T residues at the 5′-end. View largeDownload slide Figure 3. The retention times of the SN-F8-1696 oligos: (a) unmodified; (b) with six A residues at the 5′-end; (c) with six T residues at the 5′-end. References 1. Frommer,M., McDonald,L.E., Millar,D.S., Collis,C.M., Watt,F., Grigg,G.W., Molloy,P.L. and Paul,C.L. ( 1992) A genomic sequencing protocol that yields a positive display of 5-methylcytosine residues in individual DNA strands. Proc. Natl Acad. Sci. USA , 89, 1827–1831. Google Scholar 2. El-Maarri,O., Buiting,K., Peery,E.G., Kroisel,P.M., Balaban,B., Wagner,K., Urman,B., Heyd,J., Lich,C., Brannan,C.I., Walter,J. and Horsthemke.,B. ( 2001) Maternal methylation imprints on human chromosome 15 are established during or after fertilization. Nature Genet. , 3, 341–344. Google Scholar 3. Xiong,Z. and Laird,P.W. ( 1997) COBRA: a sensitive and quantitative DNA methylation assay. Nucleic Acids Res. , 12, 2532–2534. Google Scholar 4. Olek.,A., Oswald.,J. and Walter.,J. ( 1996) A modified and improved method for bisulphite based cytosine methylation analysis. Nucleic Acids Res. , 24, 5064–5066. Google Scholar 5. Gonzalgo,M.L. and Jones,P.A. ( 1997) Rapid quantitation of methylation differences at specific sites using methylation-sensitive single nucleotide primer extension (Ms-SNuPE). Nucleic Acids Res. , 12, 2529–2531. Google Scholar 6. Engemann,S., El-Maarri,O., Hajkova,P., Oswald,J. and Walter,J. ( 2001) Bisulphite-based methylation analysis of imprinted genes. In Ward,A. (ed.), Methods in Molecular Biology. Humana Press, Totowa, NJ, Vol. 181, pp. 217–228. Google Scholar 7. El-Maarri,O., Olek,A., Balaban,B., Montag,M., van der Ven,H., Urman,B., Olek,K., Caglayan,S.H., Walter,J. and Oldenburg,J. ( 1998) Methylation levels at selected CpG sites in the factor VIII and FGFR3 genes, in mature female and male germ cells: implications for male-driven evolution. Am. J. Hum. Genet. , 63, 1001–1008. Google Scholar 8. Hoogendoorn.,B., Owen.,M.J., Oefner.,P.J., Williams,N., Austin,J. and O’Donovan,M.C. ( 1999) Genotyping single nucleotide polymorphisms by primer extension and high performance liquid chromatography. Hum. Genet. , 104, 89–93. Google Scholar

Paracchini, Silvia;Arredi, Barbara;Chalk, Rod;Tyler-Smith, Chris

doi: 10.1093/nar/30.6.e27pmid: 11884646

Abstract We have established the use of a primer extension/mass spectrometry method (the PinPoint assay) for high-throughput SNP genotyping of the human Y chromosome. 118 markers were used to define 116 haplogroups and typing was organised in a hierarchical fashion. Twenty multiplex PCR/primer extension reactions were set up and each sample could be assigned to a haplogroup with only two to five of these multiplex analyses. A single aliquot of one enzyme was found to be sufficient for both PCR and primer extension. We observed 100% accuracy in blind validation tests. The technique thus provides a reliable, cost-effective and automated method for Y genotyping, and the advantages of using a hierarchical strategy can be applied to any DNA segment lacking recombination. Received November 27, 2001; Revised and Accepted January 25, 2002. INTRODUCTION Single nucleotide polymorphisms (SNPs, a term which we use to include small insertions and deletions) are the markers of choice for many applications in medical and evolutionary genetics and even forensics. They are common, widespread and stable and can cause, or be linked to, phenotypes of interest. Nevertheless, the limited number of alleles present at each variable position (usually two) often necessitates the analysis of large numbers of SNPs, sometimes in large numbers of individuals. Many SNP genotyping methods have now been developed (1). The choice of method depends on the investigation, in particular the number of SNPs and the number of individuals entering the study. Some technologies are preferred when a small number of SNPs is tested in a large population, for example the TaqMan assay (2). High density DNA arrays have proved to be a powerful tool in large-scale analyses where hundreds or thousands of SNPs are typed in a few individuals (3) and represent a promising technology, but have the disadvantage that they cannot be constructed by individual laboratories or modified to include new markers. Matrix-assisted laser desorption ionization time-of-flight (MALDI-TOF) mass spectrometry (MS) has recently been developed as a tool for SNP analysis and, although it requires a major item of equipment, several MALDI-based assays have been described (4–10). Among these methods the PinPoint assay (5) is particularly promising since it allows a high degree of multiplexing. This method is based on the addition of a single nucleotide to a genotyping primer complementary to a PCR target. The 3′-end of the genotyping primer terminates immediately upstream of the polymorphic site. Determination of the mass of the primer extension product reveals which nucleotide has been added. However, this approach has two limitations. Firstly, the mass difference between the nucleotides added can be as small as 9 Da (the difference between ddA and ddT) and, while a mass difference of this size can readily be detected, it can be difficult to distinguish between the A/T and the A/A or T/T genotypes (7). The second problem is the high level of purification required for the analysis of DNA by MALDI. We wished to establish a genotyping method for the human Y chromosome. The absence of recombination along most of its length makes it a powerful tool to study human evolution, forensics and Y-associated diseases (11). Although the discovery of Y sequence variation has been slow until recently, a large number of SNPs are now available (12). These binary markers define a well-supported and stable phylogenetic tree, which allows a simple screening strategy to be used. We designed our typing method with two features in mind. Firstly, we needed to type all of the chosen markers, so that all branches of the tree would be included: a method that typed, for example, 90% of markers would not be adequate. Secondly, it would be used to type the same set of markers in a large number of samples, so it would be worth investing time and effort in establishing an efficient method. Here we have adapted the PinPoint assay for this purpose. The problem of A-T heterozygosity does not arise since the Y chromosome is haploid; Y genotypes are thus haplotypes and are referred to here as ‘haplogroups’. The purification requirement was overcome using a commercial kit which allows sample preparation for MALDI in 96-well format. We have therefore been able to establish an efficient method and validate it by blind typing of well-characterised samples from diverse world-wide populations. MATERIALS AND METHODS Multiplex PCRs Most of the PCR primers (Table S1) were redesigned using the Primer3 program (http://www-genome.wi.mit.edu/cgi-bin/primer/primer3_www.cgi). A modified version of the protocol described by Belgrader and colleagues (13) was used for PCR. In this procedure the amplification takes place in two stages. In the first, primers consisting of a common 5′-end corresponding to zip code primer sequences of 21 or 22 bases, followed by a locus-specific sequence, are used at very low concentration. After 15 cycles, these primers are expected to be entirely incorporated into PCR products which thus have similar concentrations and the same 5′-end sequences. In the second stage, a high concentration of the zip code primers is added and the reaction is continued for another 30 cycles. In this way, even concentrations of the different products are maintained. The reaction was performed under universal conditions in a volume of 12.5 µl containing Bioline buffer [16 mM (NH4)2SO4, 67 mM Tris–HCl, pH 8.8, 0.01% Tween-20], 5 mM MgCl2, 400 µM each dNTP, ∼0.08 µM each primer, 1.25 U Biolase Diamond DNA Polymerase (Bioline) and 60 ng genomic DNA. The reaction consisted of denaturation at 94°C for 3 min, followed by 15 cycles of 94°C for 30 s, 59°C for 30 s and 72°C for 1 min, with a final extension at 72°C for 3 min. An equal volume of complete PCR buffer containing 1 µM zip code primers (ZipALg1, ggagcacgctatcccgttagac; ZipBLg2, cgctgccaactaccgcacatg) and 1.25 U enzyme was added and cycled for an additional 30 rounds as above, except that the annealing temperature was 55°C. The only modifications found to be necessary to this standard procedure were adjustments of the primer concentrations, described below, and use of the hot-start enzyme Immolase DNA Polymerase (Bioline) for multiplex PCRs numbers 11 and 15, which required an initial cycle of 7 min at 95°C. Thus, in establishing a new multiplex reaction, an initial trial would be carried out with all primers at 0.08 µM and the products examined on a gel. Primers corresponding to faint or undetectable bands were then increased in concentration and the products re-examined; this step was repeated until all bands were clearly visible and differed in molar concentration by less than 5-fold. Once established, the required primer concentrations (Table S1) were highly reproducible, except for degraded samples. Multiplex primer extensions PCR primers and dNTPs were removed by adding a ‘polishing reagent’, consisting of a phosphatase and an exonuclease (0.5 µl of Applied Biosystems 4313751 or 1 µl of Amersham Pharmacia USB 78201), to 2.5 µl of PCR products and incubation for 20 min at 37°C followed by inactivation for 20 min at 85°C. The primer extension reaction was performed by adding to the ‘polished’ PCR product a mixture containing the Biolase Diamond buffer, 5 mM MgCl2, 20 µM each ddNTP, 1 µM each genotyping primer in a final volume of 10 µl. The reaction consisted of 30 cycles of 94°C for 30 s, 37°C for 30 s and 72°C for 20 s. The genotyping primers were designed manually and are listed in Table S1. They were checked for self/cross-extension using a web-run program (http://eatworms.swmed.edu/~tim/primerfinder/) and tested experimentally in a ‘self-extension reaction’, where no PCR template was added to the primer extension mixture. Generally, no enzyme was added, but 1 U Biloase Diamond was included in the extension mix when the PCR products were obtained with Immolase DNA Polymerase. The primer extension products were desalted using a nucleic acid purification kit (Applied Biosystems 4313108) with 96-well spin plates (Applied Biosystems 4315438) following the manufacturer’s instructions. Mass spectrometry One microlitre of each sample was transferred from the microtitre plate to the inner 64 wells of a 100-well mass spectrometry plate (only these wells were fully accessible to the laser) using an eight channel Impact2 Equalizer 384 pipette (Matrix) and mixed with 1 µl of 2,4,6-trihydroxyacetophenone (THAP) (Applied Biosystems 4315786). Masses of genotyping primers and extension products were determined using a Voyager Elite DE MALDI-TOF MS workstation (PerSeptive Biosystems) in linear mode. Spectrum acquisition was completely automated, set to accumulate the first three spectra passing the acceptance criterion of a signal-to-noise ratio of 10:1. Each spectrum was derived from 35 laser shots in the same position, and after acquisition the plate was moved according to a search pattern to a different position in the same sample spot until the three spectra had been obtained. The spectra were analysed with the Data Explorer software package (PerSeptive Biosystems), using a macro that labels the primer peaks and then finds and labels the extension peaks. A macro for each multiplex was created resulting in a completely automated calling of the primers and the extension products. DNA samples Thirty samples of male genomic DNA, representing world-wide diversity (http://www.arl.arizona.edu/lmse/lmseycc.html) and previously assigned to haplogroups using other methods, were tested with this assay. Sequencing Sequencing was carried out by the sequencing service of the Department of Biochemistry, University of Oxford (http://polaris.bioch.ox.ac.uk/dnaseq/index.cfm). RESULTS Strategy Our genotyping strategy is based on the known stable phylogeny of the Y chromosome. There is no recombination on the Y chromosome outside the pseudo-autosomal regions and SNP mutation rates are low, so screening can be carried out in a hierarchical fashion, starting at the most basal branches of the tree and proceeding along the relevant branch to the tip (Fig. 1). A collection of 167 SNPs, incorporating many of those previously published, has recently been described (12), and these define 116 haplogroups. We first selected 118 markers (103 base substitutions, 11 deletions, three insertions and one complex rearrangement) which allowed all 116 haplogroups to be identified. We next assembled them into moderately sized multiplexes for both PCR amplification and primer extension. We could then place any Y chromosome into one of the 116 haplogroups using between two and five successive genotyping reactions. Genotyping procedure The genotyping procedure involves the following steps: multiplex PCR of the regions spanning the markers, enzymatic purification of the PCR products (‘polishing’), multiplex single base primer extension, DNA purification and analysis of primer extension products by MALDI-TOF MS. Twenty different multiplex reactions were designed. Each multiplex amplified between three and eight fragments and allowed the typing of between three and nine markers, because some fragments contained more than one SNP. Most locus-specific PCR primers differed from those published and were redesigned so that they would produce bands of distinguishable sizes (in the range 135–567 bp), allowing amplification to be monitored by gel electrophoresis (Fig. 2). Genotyping primers between 15 and 20 bases long were designed so that the masses of all primers and extension products in a multiplex were separated by at least 40 Da and were thus readily distinguished in the mass spectrometer. A suitable primer could be found on one or the other strand in most cases. For 11 primers it was necessary to introduce a mismatch near the 5′-end (where it would have little effect on primer binding) in order to achieve this separation, or to reduce complementarity detected in silico. When we needed to increase the separation of primers and extension products, the mass of the heavier one was increased by replacing a light base with a heavier one or, alternatively, the mass of the lighter primer was decreased. Substitutions that introduced a T were favoured because T is resistant to fragmentation during MALDI-TOF (14), e.g. C→T to increase mass and A→T or G→T to decrease mass. To reduce complementarity to no more than 3 bp between the 3′-end of a primer and a second region of any primer in the same mix, a base substitution was made in the second region, again favouring the introduction of a T. The pools were then tested experimentally for self-extension arising from self- or cross-complementarity. In two cases self-extension was seen, and for both a primer from the complementary strand was satisfactory. Primer sequences and multiplex combinations are shown in Table S1. Three aspects of the genotyping reaction were then explored: the enzymes used for PCR and primer extension, the matrix and the acquisition of data from the mass spectrometer. We tested five enzymes for PCR [Biolase Diamond DNA Polymerase (Bioline), Immolase DNA Polymerase (Bioline), AmpliTaq Gold (Applied Biosystems), AmpliTaq DNA Polymerase Stoffel Fragment (Applied Biosystems) and HotStarTaq DNA Polymerase (Qiagen)] and four for primer extension [Tth DNA Polymerase (Applied Biosystems), AmpliTaq DNA Polymerase Stoffel Fragment (Applied Biosystems), Biolase Diamond DNA Polymerase (Bioline) and HotStarTaq DNA Polymerase (Qiagen)]. Immolase, a hot-start enzyme, gave the best amplification, but Biolase Diamond DNA Polymerase was used preferentially because it provided satisfactory amplification in most cases and also performed multiplex extension. Indeed, the Biolase enzyme survived the polishing purification step between amplification and extension. It was thus unnecessary to add additional enzyme for extension, leading to a significant cost reduction. When a hot-start enzyme was used for PCR, no extension was seen with the recommended enzymes Tth DNA Polymerase (Sequazyme Pinpoint SNP Assay Kit; Applied Biosystems) and DNA Polymerase Stoffel Fragment (5), but extension was obtained with Biolase Diamond DNA Polymerase. Two MALDI matrices were compared: THAP and 3-hydroxypicolinic acid. THAP was preferred because it gave more even crystallization, increasing the data acquisition speed in automated mode, as also reported elsewhere (15). Different mass spectrometer data acquisition modes were investigated. Each spectrum takes ∼12 s to acquire and over 100 spectra can be generated from different positions in the same sample spot. The mass spectrometer was therefore set to accumulate and save the first three spectra that passed our chosen acceptance criterion. This took 1 min/sample, on average. Validation All 20 pools showed the expected extension products. Typical results are presented in Figure 3. To test the accuracy of this assay, we carried out blind typing of 30 diverse samples which had previously been genotyped with the entire set of markers by other methods. These individuals required between two and four multiplex reactions each and used 15 of the 20 multiplexes, requiring just 78 PCRs/extensions in all. Samples were assigned to 14 different haplogroups. Twenty-nine out of the 30 samples gave the expected results, but one showed a single discrepancy at a single locus, where our typing revealed the derived C allele instead of the expected ancestral G allele at the M35 locus [according to the terminology of Underhill et al. (12), which uses the opposite strand from Table S1 for this locus]. Sequencing showed that a C was indeed present at this position (S.Paracchini, unpublished observations) and thus that the discrepancy was a result of previous mistyping or sample mix-up; the accuracy of the MS method was 100%. The complete set of 3540 SNPs (118 in 30 chromosomes) was evaluated, but only 492 needed to be tested experimentally because of our hierarchical strategy. DISCUSSION We have established a quick and efficient method for SNP genotyping of the human Y chromosome which places samples in one of 116 haplogroups after between two and five multiplex reactions. All 118 markers investigated could be typed by this method, although for 2/118 the first genotyping primer tested was unsatisfactory because of extension in the absence of template due to self-complementarity, and it was necessary to use the primer from the complementary strand. The proportion of SNPs that cannot be typed at all because both primers are unsatisfactory is likely to be very low. Our typing strategy maximises efficiency by its hierarchical organisation, but does this sacrifice information? Recurrent mutations on branches that are not typed will be missed, but haplogroup assignment will always be correct. Such recurrent mutations are rare and generally the large benefits of saving DNA, time and expense will outweigh any small loss of information. For samples where typing of all 118 markers is required, this can still be achieved in 20 reactions. The use of multiplexes has the additional advantage of reducing the risk of errors in associating SNP allelic states into haplogroups, and the automated calling of the peaks further reduces opportunities for human error. Typing is thus very accurate. Haplogroup assignment, implying correct typing of all 118 markers, was 100% correct in all samples tested and actually revealed an error in the previous data. Several factors contribute to this accuracy. Failure to amplify or extend produces only the primer peak in the mass spectrum and cannot be misinterpreted as the presence of one of the alleles. Spurious peaks are sometimes generated by depurination (Fig. 3A and B, asterisks), but can all be distinguished from primers and extension products by their mass. Mixed samples are sometimes encountered, usually as a result of unintentional sample contamination, and these can readily be recognised. Heterozygotes, where A and T extensions can be difficult to distinguish, do not occur on the Y chromosome. Nevertheless, duplicated Y loci can potentially produce two extension products from a single primer, which might lead to similar confusion. Duplicated loci were rare among the Y markers used and none involved the addition of an A or a T to the same primer. If there was such a case, a mass-tagged ddA analogue could be used (7). Potential A/T confusion is avoided in different ways by other MALDI-TOF methods (4,10), but it is unclear whether a high degree of multiplexing can be achieved using such methods. Throughput is high: at 1 min/sample, an average of six SNPs per multiplex and automated sample spotting and MS plate changing, more than 8000 SNPs/day could be typed. The major expenses are the enzymes and purification kit; we estimate that the consumables cost per SNP was ∼30 p (∼US$ 0.4) under the conditions used. This could be reduced to ∼17 p (∼US$ 0.24) per SNP by reducing the PCR volume to 2.5 µl (the minimum required) and re-using the 96-well spin plates. These costs compare favourably with other methods. All PCRs are carried out under the same standard conditions and we have found that additional loci can be incorporated into the multiplex reactions, making it flexible enough to accommodate new markers that may be discovered in the future. Although access to a MALDI-TOF mass spectrometer is required, our strategy of hierarchical multiplex PCR and primer extension reactions can readily be adapted for other platforms, for example by using fluorescently labelled ddNTPs. A hierarchical screening strategy can be applied to any non-recombining DNA segment. While this would seem to limit it to the Y chromosome and mitochondrial DNA in humans, it has recently been suggested that recombination in autosomal DNA is largely or entirely confined to small ‘hot-spots’, so that most SNPs lie in discrete segments where there is little or no recombination (16). If this is true, each segment could be screened in a hierarchical fashion. In our analysis complete information for 30 individuals with 118 markers (3450 SNPs) was obtained by typing only 492 SNPs and inferring the state at the remainder: a 7-fold reduction. In a large-scale genotyping project this would produce significant savings. We conclude that this method provides a high-throughput assay to genotype the Y chromosome that is accurate, robust, reliable, flexible, cost-effective and lends itself to automation. SUPPLEMENTARY MATERIAL Supplementary Material is available at NAR Online. ACKNOWLEDGEMENTS We thank Ed Southern for discussions and comments on the manuscript, M. Sohail for advice and Martin Johnson for maintaining the mass spectrometer. S.P and C.T.-S. were supported by the CRC. B.A. was supported by the Immunohematology Laboratory, Istituto di Medicina Legale, Università Cattolica del Sacro Cuore, Rome, Italy. * To whom correspondence should be addressed. Tel: +44 1865 275222; Fax: +44 1865 275259; Email: [email protected] Present address:Rod Chalk, Oxford GlycoSciences (UK) Ltd, 86 Milton Park, Abingdon, Oxon OX14 4RY, UK View largeDownload slide Figure 1. Hierarchical screening strategy. The tree shows the Y chromosomal phylogeny defined by 167 markers, represented here by 118 SNPs (numbers on tree) identifying 116 haplogroups (italic numbers on the right side). The SNPs included in each multiplex are grouped in a grey area marked with the multiplex number (large bold type) from Table S1. View largeDownload slide Figure 1. Hierarchical screening strategy. The tree shows the Y chromosomal phylogeny defined by 167 markers, represented here by 118 SNPs (numbers on tree) identifying 116 haplogroups (italic numbers on the right side). The SNPs included in each multiplex are grouped in a grey area marked with the multiplex number (large bold type) from Table S1. View largeDownload slide Figure 2. Multiplex PCR products. Products generated with multiplex 9 from three individuals (tracks 1–3) or a sample lacking DNA (Blank) were analysed by electrophoresis on a 3% agarose gel and visualised by ethidium bromide staining. Eight bands are seen, as expected (right side). View largeDownload slide Figure 2. Multiplex PCR products. Products generated with multiplex 9 from three individuals (tracks 1–3) or a sample lacking DNA (Blank) were analysed by electrophoresis on a 3% agarose gel and visualised by ethidium bromide staining. Eight bands are seen, as expected (right side). View largeDownload slide Figure 3. Multiplex primer extension products analysed by mass spectrometry. (A) Multiplex 1, individual 1. (B) Multiplex 1, individual 2. (C) Multiplex 4, individual 3. Automatic calling of primers (-P) or SNPs (-A, -C, -G, -T) is shown within the body of each section and the expected positions are shown at the top. Asterisks indicate prominent depurination products. The allelic differences for the same multiplex between individuals 1 and 2 can be seen. View largeDownload slide Figure 3. Multiplex primer extension products analysed by mass spectrometry. (A) Multiplex 1, individual 1. (B) Multiplex 1, individual 2. (C) Multiplex 4, individual 3. Automatic calling of primers (-P) or SNPs (-A, -C, -G, -T) is shown within the body of each section and the expected positions are shown at the top. Asterisks indicate prominent depurination products. The allelic differences for the same multiplex between individuals 1 and 2 can be seen. References 1. Gut,I.G. 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Ryder, Sean P.;Strobel, Scott A.

doi: 10.1093/nar/30.6.1287pmid: 11884625

Abstract Great strides in understanding the molecular underpinnings of RNA catalysis have been achieved with advances in RNA structure determination by NMR spectroscopy and X-ray crystallography. Despite these successes the functional relevance of a given structure can only be assessed upon comparison with biochemical studies performed on functioning RNA molecules. The hairpin ribozyme presents an excellent case study for such a comparison. The active site is comprised of two stems each with an internal loop that forms a series of non-canonical base pairs. These loops dock into each other to create an active site for catalysis. Recently, three independent structures have been determined for this catalytic RNA, including two NMR structures of the isolated loop A and loop B stems and a high-resolution crystal structure of both loops in a docked conformation. These structures differ significantly both in their tertiary fold and the nature of the non-canonical base pairs formed within each loop. Several of the chemical groups required to achieve a functioning hairpin ribozyme have been determined by nucleotide analog interference mapping (NAIM). Here we compare the three hairpin structures with previously published NAIM data to assess the convergence between the structural and functional data. While there is significant disparity between the interference data and the individual NMR loop structures, there is almost complete congruity with the X-ray structure. The only significant differences cluster around an occluded pocket adjacent to the scissile phosphate. These local differences may suggest a role for these atoms in the transition state, either directly in chemistry or via a local structural rearrangement. Received November 16, 2001; Revised and Accepted January 14, 2002. INTRODUCTION Nucleotide analog interference mapping (NAIM) is an efficient chemical group mutagenesis strategy used to identify the subset of atoms important for the function of an RNA molecule (1,2). The method identifies chemical groups involved in secondary or tertiary hydrogen bonds, ligand binding, active site chemistry or other roles required for RNA activity. These data should be useful for analyzing the functional relevance of three-dimensional RNA structures because chemical groups that display an effect on RNA activity when modified or deleted should be the same groups structurally implicated for their involvement in activity (3,4). If there is only a weak correlation between the structural and biochemical data, then it suggests that the interactions observed in the structure are not relevant to the functioning RNA or that the structure does not represent the active state. In either case, NAIM analysis is highly complementary to structure determination, as it can ascribe functional relevance to contacts observed within a structure. The intent of this report is to compare previously published interference data with recent structures obtained for the hairpin ribozyme, a small catalytic RNA motif that performs a reversible self-cleavage reaction. In nature, the ribozyme functions in the processing of replication intermediates in the ‘life’ cycle of a tobacco ringspot nepovirus satellite RNA (5). In this context, the ribozyme folds into an extended cruciform structure (Fig. 1) (6). The minimum elements of the hairpin RNA required for catalytic activity in vitro are just two arms of this cruciform, which include four short helical elements (helix 1–4) and two internal loop regions, A and B (7,8). The ribozyme performs a cleavage reaction at a specific site within the symmetric loop A, yielding two products with a 5′‐hydroxyl group and a 2′,3′-cyclic phosphate, respectively (5). The products of this reaction are similar to three other naturally occurring catalytic RNAs, the hammerhead, VS and HDV ribozymes, although significant differences in secondary structure imply that each molecule has its own distinct catalytic strategy for arriving at these products (9). Three structures have been reported for the hairpin ribozyme. These include two NMR structures of the isolated stems containing loop A and loop B (10,11) and a high-resolution crystal structure of the docked complex with a 2′-O-methyl inhibitor in place of the 2′-OH nucleophile (12). These structures differ significantly in both their tertiary structure fold and in the nature of the non-canonical base pairs formed in each loop. Such dramatic differences raise a significant question: which if any of these structures represent the hairpin ribozyme in its active conformation? Previously, we performed NAIM analysis on the hairpin with several nucleotide analogs of A, C, G and U (13). These included analogs that eliminated or modified functional groups on either the heterocyclic base or the ribose sugar. Here, we utilize these data to assess the functional relevance of the available hairpin ribozyme structures. HAIRPIN RIBOZYME STRUCTURES Two independent NMR structures have been determined for the hairpin ribozyme. The structure of helix 1, loop A, and helix 2 (termed stem A) was solved in isolation from the rest of the ribozyme by Cai and Tinoco (11) (Fig. 2). In this structure, the bases of loop A form a series of four non-canonical pairs resulting in an extended helix. One base, U+2, flips out of the helix and is oriented toward the solvent. The other half of the ribozyme was also determined by NMR. Butcher and coworkers (10) reported the structure of helix 3, loop B, and helix 4 (termed stem B) (Fig. 3). Similar to the loop A structure, the nucleotides in loop B form a series of non-canonical base pairs with two bulged uridines, U39 and U41. The geometry of the non-canonical pairing disrupts the continuous stack, so that helix 3 stacks upon four pairs in the top of loop B, while helix 4 stacks upon the three pairs at the bottom of loop B. As a result, this stem adopts a bent structure with a narrow minor groove that appears to have some conformational flexibility. Though stem A and stem B each form an independently folded unit, the authors suggest that conformational changes may be necessary to achieve the active, docked structure. Rupert and Ferre-D’Amare recently reported the 2.4 Å structure of the intact ribozyme in which loop A is docked into loop B (Fig. 4) (12). In this context the secondary structures in loop A and loop B are significantly different than in the isolated stems. For example, all of the non-canonical base pairs observed in the NMR structure of stem A adopt a different conformation in the docked structure. Instead of being flipped out of the helix, U+2 is in the helical stack and pairs with G8, while G+1 adopts an unusual 2′-endo sugar pucker and syn base configuration and base pairs with C25 in loop B (12,14). Likewise, several differences are observed in the non-canonical base pairs observed in stem B. These changes are necessary to facilitate specific tertiary interactions between loops A and B. SUPERPOSITION OF NAIM DATA ON THE HAIRPIN STRUCTURES In an effort to systematically compare the data obtained on active hairpin ribozymes with the available hairpin structures, we superimposed the interference data collected with all of the analogs onto the coordinates of each structure (Figs 2–4). The interferences used in this comparison are listed in Table 1. We used the following criterion to assess the correlation between each structure and the biochemical data. (i) A given interference is considered to be agreement if the analog deletes a functional group involved in an interaction within the structure. For example, the N2 amine of G21 interacts with the N7 of A22 in the loop B NMR structure. Both of these groups lead to interference when modified, hence this contact is consistent with the interference data. (ii) An interference resulting from incorporation of a bulky nucleotide analog matches the structure if it creates a steric clash, which implies that the structure cannot sufficiently accommodate the analog modification. For example, the N2 amine of G8 makes two contacts via its N2 amine in the crystal structure to the 2′-OH of A-1 and the O2 of U+2. Consistent with these contacts, methylation of this amine disrupts activity. (iii) An analog may affect the local conformation of the nucleotide, for example disruption of the sugar pucker. An example of this occurs at U41 with interference from 2′‐fluoro substitution, an analog that favors a C-3′-endo conformation. U41 base adopts C-2′-endo in both the NMR and the crystal structure and hence 2′-fluoro interference is consistent with both structures. For interferences that meet these criteria, the affected functional groups are designated with blue colored spheres in Figures 2–4. If none of these criteria are met within the structure, the functional groups that show interference are designated with yellow colored spheres. STEM A AND B STRUCTURES ARE LARGELY INCONSISTENT WITH THE NAIM DATA Even a cursory inspection of the figures shows that the isolated stem A and stem B NMR structures are not well matched with the NAIM data (Figs 2 and 3). In the structure of loop A, most of the interferences (>50%) cannot be explained by the structure. Similarly, the loop B NMR structure does not match well with the interference pattern. It is possible that the failure of some functional groups to appear relevant within the NMR structures could result from a lack of tertiary contacts to the opposing stem in the functional docked structure. For example, several interferences at positions A9-U12 do not reflect contacts within the secondary structure of the stem, but they cluster in the minor groove surface of the loop A secondary structure. This may reflect a pre-ordered loop B binding surface. However, there is reason to doubt that the stem A NMR structure represents a pre-folded structural domain that utilizes this minor groove surface to interact with stem B in the functioning ribozyme. Several functional groups expected to be necessary for ribozyme activity based on the geometry of stem A fail to show interference with analogs that delete or modify them. While these are negative data, they may indicate that certain aspects of the NMR structure do not adequately reflect the docked, active structure. For example, the N2 exocyclic amine of G+1 was shown to be critical for ribozyme activity (15,16). Site-specific inosine substitution at this position reduces ribozyme cleavage activity to below detectable levels. In the NMR structure, this amine interacts with the N7 of A9. Analogs that delete or modify the N7 moiety do not interfere with ribozyme activity at A9. If the sole role of G+1 amine is to form a single H-bond interaction with the A9 N7, then analogs that disrupt this position should affect ribozyme activity at a level comparable with the G+1 inosine effect. While it is possible that the G+1 amine may form an additional contact in the active structure, these data argue that the stem A structure might not reflect that conformation. Unlike the interferences in stem A, those in stem B do not cluster into a discreet region of the structure. Instead, they are scattered across the minor and major grooves throughout the length of the molecule. As such, these interferences are not likely to reflect the formation of tertiary contacts missing in the isolated loop B stem structure. It is more likely that the secondary structure of loop B undergoes a rearrangement prior to forming the docked structure. THE CRYSTAL STRUCTURE MATCHES THE NAIM DATA EXCEPT NEAR THE SCISSILE PHOSPHATE In contrast to the isolated stem structures, the crystal structure of the intact ribozyme is almost fully consistent with the NAIM data (Fig. 4). Approximately 85% of the interferences can be explained by features in this structure. Interference is observed at most of the functional groups involved in the non-canonical secondary structure in stems A and B, as well as those in tertiary interface. Bulky analogs are accommodated on the surface of this structure, but not in the tightly packed interface between the internal loops. All of the unexplained interferences cluster around the scissile phosphate in a pocket formed by the conserved residues G+1, A9, A10, C25 and A38 (12). All of the functional groups that point into this cavity are required for catalysis, including the N2 amine of G+1, the N6 amine of A38, and the N1 imino group of A10 (2,13,15). Furthermore, occupation of this cavity by methylation of the N6 amine of A9 significantly disrupts activity (13). Previous work indicated that ionization of A10 is important for ribozyme catalysis, that the N6 amine of A38 is indispensable for catalysis, and that these groups are likely to be involved in the stabilization of the catalytic transition state (17). These few exceptions may reflect local disturbances in the structure necessary to accommodate the 2′-O-methyl substitution at the cleavage site or transition state specific effects on ribozyme chemistry. They may provide clues into the mechanism of catalysis by the hairpin ribozyme. INTERPRETATION AND ANALYSIS Several lines of evidence suggest that the crystal structure of the docked complex more accurately reflects the biologically relevant structure than either NMR structure. (i) The strong correlation between the NAIM data and the crystal structure evident from Figure 4 in this analysis. (ii) The nucleophile, reactive phosphate, and 5′-oxy leaving group are in an in-line configuration (12). This specific geometry is necessary for the SN2 phosphodiester reaction, but is not the geometry of the scissile phosphate within the loop A structure (11). (iii) The crystal structure matches the artificial phylogenetic data on the ribozyme. All of the conserved nucleotides in the hairpin sequence are involved in specific secondary and tertiary structure contacts (18). (iv) Several specific tertiary interactions predicted by crosslinking and mutagenesis experiments are observed in the crystal structure (8,14,19). (v) The structure is consistent with the majority of solution modification experiments and site-specific analog substitution data (8,16,20–24). (vi) Hampel and Burke have shown that a highly reactive photo-crosslink indicative of the ‘loop E-like’ motif present in the loop B NMR structure is inhibitory to docking and ribozyme catalysis and that the crosslink is lost upon formation of the active structure (21). Although the biochemical data suggest that the NMR structures of stem A and stem B do not reflect the active conformation, it does not mean that these structures are incorrect. It is likely that each structure reflects the accurate, undocked conformation. In fact, this interpretation is suggested by crosslinks observed in stem B that are indicative of features observed in the NMR structure (21). Furthermore, the lack of observable imino NOEs for the loop region of stem B, as well as intermediate J-couplings for the majority of the bases, suggest that the structure of this molecule is unusually dynamic (10). If true, the hairpin ribozyme must undergo a complex series of rearrangements in order to form the docked complex. This is corroborated by FRET studies that suggest docking limits the rate of ligation chemistry (25). While the NAIM data strongly support the functional relevance of the crystal structure, a handful of the most interesting interferences cluster in a cavity surrounding the scissile phosphate and are not explained by the structure. These groups do not appear to be making ground state contacts within the tertiary structure, but instead may participate in promoting the chemical transition. One possibility is that the cavity is occupied by a reaction molecule, such as a water, and that these groups are in position to coordinate and activate it for catalysis. While no water molecule was observed in the cavity within the hairpin crystal structure, the authors note that there is sufficient room for one to be accommodated (12). Alternately, ionization of A10 could stabilize charge in the pre-ligation ground state or in the transition state by interacting with a non-bridging phosphate oxygen of the scissile phosphate, while a subtle change in geometry could allow the amine of A38 to hydrogen bond to the 5′-hydroxyl nucleophile in the ligation reaction. Although this model would require alterations in the geometry of the bases that surround the reactive phosphate, it is possible that removal of the 2′-O-methyl inhibitor or subtle differences between the cleaved and ligated forms of the ribozyme could lead to these changes. In either case, movement of A10 and A38 by <3 Å would position these bases for a direct role in catalysis. In summary, the NAIM data argue that the docked crystal structure very closely reflects the functioning state of the hairpin ribozyme. However, the crystal structure does not immediately elucidate the mechanism of hairpin ribozyme catalysis. Strikingly, the only positions of interference that do not correlate to the crystal structure lie in the vicinity of the active site. It is possible that these groups are involved in hairpin ribozyme catalysis via an as yet to be determined chemical pathway. ACKNOWLEDGEMENTS We would like to thank Michael Recht and Pete Funke for assistance with preparing figures. This work was supported by NSF grant CHE-0100057 to S.A.S. * To whom correspondence should be addressed. Tel: +1 203 432 9772; Fax: +1 203 432 5767; Email: [email protected] Present address:Sean P. Ryder, The Scripps Research Institute, Department of Molecular Biology, 10550 North Torrey Pines Road, MB-33, La Jolla, CA 93027, USA View largeDownload slide Figure 1. Schematic secondary structure of the hairpin ribozyme in its natural context. The hairpin ribozyme motif is in bold line. The remainder of the satellite RNA sequence is represented by dashed lines. The four stems of the ribozyme are labeled. The cleavage site is marked with an arrow. View largeDownload slide Figure 1. Schematic secondary structure of the hairpin ribozyme in its natural context. The hairpin ribozyme motif is in bold line. The remainder of the satellite RNA sequence is represented by dashed lines. The four stems of the ribozyme are labeled. The cleavage site is marked with an arrow. View largeDownload slide Figure 2. NAIM compared with the stem A NMR structure. Sites of interference are denoted by spheres. Blue spheres represent sites of interference consistent with the structure. The yellow spheres represent sites that cannot be interpreted in comparison with the structure. The scissile phosphate is denoted by a red sphere. View largeDownload slide Figure 2. NAIM compared with the stem A NMR structure. Sites of interference are denoted by spheres. Blue spheres represent sites of interference consistent with the structure. The yellow spheres represent sites that cannot be interpreted in comparison with the structure. The scissile phosphate is denoted by a red sphere. View largeDownload slide Figure 3. NAIM compared with the NMR structure of stem B. The diagram is labeled as in Figure 2. View largeDownload slide Figure 3. NAIM compared with the NMR structure of stem B. The diagram is labeled as in Figure 2. View largeDownload slide Figure 4. NAIM compared with the crystal structure of the docked ribozyme. The diagram is labeled as in Figure 2. View largeDownload slide Figure 4. NAIM compared with the crystal structure of the docked ribozyme. The diagram is labeled as in Figure 2. Table 1. NAIM analysis of the hairpin ribozyme Position  Analog effects  Comment  G8  Ino, m2G    A9  m6A, DAP, Pur, 2AP, c3A, n8A  n8A is an enhancement, attributed to the N1 position (17)  A10  Pur, 2AP, dA, FA, c3A, n8A  n8A interference is pH dependent, attributed to N1 ionization (17)  G11  Ino, m2G, dG, FG    U12  dU, FU    G21  Ino, m2G    A20  m6A    A22  7dA, m6A    A23  m6A, DAP, 2AP    A24  DAP, Pur, FA, c3A    C25  dC, 5mFC, Zeb    A26  DAP, 2AP    A38  7dA, DAP, Pur, 2AP, dA, FA    U39  FU, m5U    A40  DAP, 2AP, c3A    U41  FU    U42  5mU, dU  dU is an enhancement, attributed to increased conformational flexibility (13)  A43  7dA    C44  Zeb    Position  Analog effects  Comment  G8  Ino, m2G    A9  m6A, DAP, Pur, 2AP, c3A, n8A  n8A is an enhancement, attributed to the N1 position (17)  A10  Pur, 2AP, dA, FA, c3A, n8A  n8A interference is pH dependent, attributed to N1 ionization (17)  G11  Ino, m2G, dG, FG    U12  dU, FU    G21  Ino, m2G    A20  m6A    A22  7dA, m6A    A23  m6A, DAP, 2AP    A24  DAP, Pur, FA, c3A    C25  dC, 5mFC, Zeb    A26  DAP, 2AP    A38  7dA, DAP, Pur, 2AP, dA, FA    U39  FU, m5U    A40  DAP, 2AP, c3A    U41  FU    U42  5mU, dU  dU is an enhancement, attributed to increased conformational flexibility (13)  A43  7dA    C44  Zeb    Ino, inosine; m2G, N2-methylguanosine; dG, 2′-deoxyguanosine; FG, 2′-deoxy-2′-fluoroguanosine; m6A, N6-methyladenosine; DAP, 2,6-diaminopurine riboside; Pur, purine riboside; 2AP, 2-aminopurine riboside; c3A, 3-deazaadenosine; n8A, 8-azaadenosine; dA, 2′-deoxyadenosine; FA, 2′-deoxy-2′-fluoroadenosine, 7dA, 7-deazaadenosine; dC, 2′-deoxycytidine; 5mFC, 5-methyl-2′-fluorocytidine; Zeb, Zebularine; dU, 2′-deoxyuridine; FU, 2′-deoxy-2′-fluorouridine; 5mU, 5-methyluridine (ribothymidine). 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Mackay, Ian M.;Arden, Katherine E.;Nitsche, Andreas

doi: 10.1093/nar/30.6.1292pmid: 11884626

Abstract The use of the polymerase chain reaction (PCR) in molecular diagnostics has increased to the point where it is now accepted as the gold standard for detecting nucleic acids from a number of origins and it has become an essential tool in the research laboratory. Real-time PCR has engendered wider acceptance of the PCR due to its improved rapidity, sensitivity, reproducibility and the reduced risk of carry-over contamination. There are currently five main chemistries used for the detection of PCR product during real-time PCR. These are the DNA binding fluorophores, the 5′ endonuclease, adjacent linear and hairpin oligoprobes and the self-fluorescing amplicons, which are described in detail. We also discuss factors that have restricted the development of multiplex real-time PCR as well as the role of real-time PCR in quantitating nucleic acids. Both amplification hardware and the fluorogenic detection chemistries have evolved rapidly as the understanding of real-time PCR has developed and this review aims to update the scientist on the current state of the art. We describe the background, advantages and limitations of real-time PCR and we review the literature as it applies to virus detection in the routine and research laboratory in order to focus on one of the many areas in which the application of real-time PCR has provided significant methodological benefits and improved patient outcomes. However, the technology discussed has been applied to other areas of microbiology as well as studies of gene expression and genetic disease. Received October 31, 2001; Revised January 2, 2002; Accepted January 14, 2002. BACKGROUND The polymerase chain reaction (PCR) (1,2) has been used as the new gold standard for detecting a wide variety of templates across a range of scientific specialties, including virology. The method utilises a pair of synthetic oligonucleotides or primers, each hybridising to one strand of a double-stranded DNA (dsDNA) target, with the pair spanning a region that will be exponentially reproduced. The hybridised primer acts as a substrate for a DNA polymerase (most commonly derived from the thermophilic bacterium Thermus aquaticus and called Taq), which creates a complementary strand via sequential addition of deoxynucleotides. The process can be summarised in three steps: (i) dsDNA separation at temperatures >90°C, (ii) primer annealing at 50–75°C, and (iii) optimal extension at 72–78°C (Fig. 1A). The rate of temperature change or ramp rate, the length of the incubation at each temperature and the number of times each set of temperatures (or cycle) is repeated are controlled by a programmable thermal cycler. Current technologies have significantly shortened the ramp times using electronically controlled heating blocks or fan-forced heated air flows to moderate the reaction temperature. Consequently, PCR is displacing some of the gold standard cell culture, antigenaemia and serological assays (3). Existing combinations of PCR and detection assays (called ‘conventional PCR’ here) have been used to obtain quantitative data with promising results. However, these approaches have suffered from the laborious post-PCR handling steps required to evaluate the amplicon (4). Traditional detection of amplified DNA relies upon electrophoresis of the nucleic acids in the presence of ethidium bromide and visual or densitometric analysis of the resulting bands after irradiation by ultraviolet light (5). Southern blot detection of amplicon using hybridisation with a labelled oligonucleotide probe is also time consuming and requires multiple PCR product handling steps, further risking a spread of amplicon throughout the laboratory (6). Alternatively, PCR–ELISA may be used to capture amplicon onto a solid phase using biotin or digoxigenin-labelled primers, oligonucleotide probes (oligoprobes) or directly after incorporation of the digoxigenin into the amplicon (7–12). Once captured, the amplicon can be detected using an enzyme-labelled avidin or anti-digoxigenin reporter molecule similar to a standard ELISA format. The possibility that, in contrast to conventional assays, the detection of amplicon could be visualised as the amplification progressed was a welcome one (13). This approach has provided a great deal of insight into the kinetics of the reaction and it is the foundation of kinetic or ‘real-time’ PCR (Fig. 1B) (6,14–17). Real-time PCR has already proven itself valuable in laboratories around the globe, building on the enormous amount of data generated by conventional PCR assays. The monitoring of accumulating amplicon in real time has been made possible by the labelling of primers, probes or amplicon with fluorogenic molecules, This chemistry has clear benefits over radiogenic oligoprobes that include an avoidance of radioactive emissions, ease of disposal and an extended shelf life (18). The increased speed of real-time PCR is largely due to reduced cycle times, removal of post-PCR detection procedures and the use of fluorogenic labels and sensitive methods of detecting their emissions (19,20). The reduction in amplicon size generally recommended by the creators of commercial real-time assays may also play a role in this speed, however we have shown that decreased product size does not necessarily improve PCR efficiency (21). The disadvantages of using real-time PCR in comparison with conventional PCR include the inability to monitor amplicon size without opening the system, the incompatibility of some platforms with some fluorogenic chemistries, and the relatively restricted multiplex capabilities of current applications. Also, the start-up expense of real-time PCR may be prohibitive when used in low-throughput laboratories. These shortcomings are mostly due to limitations in the system hardware or the available fluorogenic dyes or ‘fluorophores’, both of which will be discussed in more detail. Because most of the popular real-time PCR chemistries depend upon the hybridisation of an oligoprobe to its complementary sequence on one of the strands of the amplicon, the use of more of the primer that creates this strand is beneficial to the generation of an increased fluorescent signal (22). Asymmetric PCR, as this is known, has been shown to produce improved fluorescence from a hairpin oligoprobe PCR (23) and we have found it directly applicable to other oligoprobe-hybridisation assays. The most commonly used fluorogenic oligoprobes rely upon fluorescence resonance energy transfer (FRET; Fig. 2) between fluorogenic labels or between one fluorophore and a dark or ‘black-hole’ non-fluorescent quencher (NFQ), which disperses energy as heat rather than fluorescence. FRET is a spectroscopic process by which energy is passed between molecules separated by 10–100 Å that have overlapping emission and absorption spectra (24,25). Förster primarily developed the theory behind this process: the mechanism is a non-radiative induced-dipole interaction (26). The efficiency of energy transfer is proportional to the inverse sixth power of the distance (R) between the donor and acceptor (1/R6) fluorophores (27,28). Post-amplification manipulation of the amplicon is not required for real-time PCR, therefore these assays are described as ‘closed’ or homogeneous systems. The advantages of homogeneous systems include a reduced result turnaround, minimisation of the potential for carry-over contamination and the ability to closely scrutinise the assay’s performance (29). In addition, real-time PCR has proven cost effective when implemented in a high throughput laboratory (30), particularly when replacing conventional, culture-based approaches to virus detection. In the remainder of this article, the theory behind real-time PCR will be reviewed and its rapidly increasing use in the fields of human virology will be used as an illustration. AMPLICON DETECTION In the following section we will focus on the detection processes that discriminate real-time PCR from conventional PCR assays. There are five major chemistries currently in use, and they can be classified into amplicon sequence specific or non-specific methods of real-time PCR detection (31). Each of the chemistries has an associated nomenclature to describe the fluorescent labels; however, for general discussion, fluorophore will continue to be used to describe these moieties. Although this review focuses on the use of these chemistries in real-time applications, they can also be used as a label for end-point amplicon detection. DNA-binding fluorophores The basis of the sequence non-specific detection methods is the DNA-binding fluorogenic molecule. Included in this group are the earliest and simplest approaches to real-time PCR. Ethidium bromide (32), YO-PRO-1 (33,34) and SYBR® green 1 (35) all fluoresce when associated with dsDNA which is exposed to a suitable wavelength of light. This approach requires less specialist knowledge than the design of fluorogenic oligoprobes, is less expensive and does not suffer when the template sequence varies, which may abrogate hybridisation of an oligoprobe (36). Formation of primer-dimer (37) is common and, together with the formation of specific products, is strongly associated with entry of the PCR into the plateau phase (Fig. 1B) (38,39). Association of a DNA-binding fluorophore with primer-dimer or other non-specific amplification products can confuse interpretation of the results. Adding a short, higher temperature incubation after the extension step in which fluorescence data are acquired minimises the contribution of these products to the fluorescence signal (40). The problem of primer-dimer can also be addressed using software capable of fluorescent melting curve analysis. This method makes use of the temperature at which the dsDNA amplicon is denatured (TD; Fig. 1A). The shorter primer-dimer can be discriminated by its reduced TD compared with the full-length amplicon. Analysis of the melting curves of amplicon in the presence of SYBR green 1 has demonstrated that the practical sensitivity of DNA-binding fluorophores is limited by non-specific amplification at low initial template concentrations. DNA binding fluorophores also increase the TD and broaden the melting transition, requiring substantial sequence change to produce a shift in the TD. Oligoprobes are able to discriminate single point mutations using the temperature at which 50% of oligoprobe-target duplexes separate (41). This temperature is called the melting temperature (TM) and it is dependent upon the concentration of the dsDNA, its length, nucleotide sequence and the solvent composition, and is often confused with TD (Fig. 1A) (42). Linear oligoprobes The use of a pair of adjacent, fluorogenic hybridisation oligoprobes was first described in the late 1980s (43,44) and, now known as ‘HybProbes’, they have become the method of choice for the LightCycler™ (Roche Molecular Biochemicals, Germany), a capillary-based, microvolume fluorimeter and thermocycler with rapid temperature control (20,45). The upstream oligoprobe is labelled with a 3′ donor fluorophore (FITC) and the downstream probe is commonly labelled with either a LightCycler Red 640 or Red 705 acceptor fluorophore at the 5′ terminus so that when both oligoprobes are hybridised, the two fluorophores are located within 10 nt of each other, sometimes attracting the name ‘kissing’ probes (Figs 2C, D and 3C). The plastic and glass composite capillaries are optically clear and act as cuvettes for fluorescence analysis, as well as facilitating rapid heat transfer. Capillaries are rotated past a blue light-emitting diode and fluorescence is monitored by three photodetection diodes with different wavelength filters. The temperature is varied by rapidly heating and cooling air using a heating element and fan which produce ramp rates of 20°C/s, prolonging polymerase survival (46). Additionally, because the oligoprobes are not significantly hydrolysed during amplification (47) and the LightCycler is able to monitor the changes in fluorescence emission during denaturation of the adjacent oligoprobes from their amplicon, this system can perform single tube genotyping. This capability, which makes use of fluorescent melting curve analysis, provides significant information about the sequence to which the oligoprobes are binding. Mutation(s) under one or both oligoprobes can be determined by the decrease in melting temperature that they incur due to destabilisation of the oligoprobe/target duplex (Fig. 4). This has imparted significant improvements in speed upon the diagnosis of genetic disease as well as a growing number of multiplex PCR approaches for the detection of related viral pathogens. Despite the fact that the hybridisation does not reach equilibrium using these ramp rates, the apparent TM values are both reproducible and characteristic of a given probe/target duplex (48). However, the capillaries are fragile and require some experience to handle (49). When comparing signals from the different chemistries, the destruction of nuclease oligoprobes continues despite a plateau in product accumulation whereas SYBR green 1 fluorescence in the no template control generally increases non-specifically during later cycles. Adjacent oligoprobe fluorescence begins to decrease as the rate of collision between the growing numbers of complementary amplicon strands increases favouring the formation of dsDNA over the hybridisation of oligoprobe to its target DNA strand. Additionally, there is the possibility that some oligoprobe is consumed by sequence-related endonuclease activity (50,51). All three oligoprobe chemistries (SYBR Green I, nuclease and adjacent oligoprobes) seem capable of detecting amplified product with approximately the same sensitivity (20). Combinations of the above approaches are now appearing as more users of the instrumentation become familiar with the concepts behind real-time PCR and contribute to the literature. If a sequence-specific, fluorophore-labelled linear oligoprobe is added to a SYBR green 1 mix, currently called the Bi-probe system, FRET will occur and an additional layer of specificity can be obtained (44,52,53). An assay using a BODIPY®FL-labelled oligoprobe was adapted to run in the LightCycler using a β-globin target sequence (54). The probe was designed so that the fluorophore was located on a terminal cytosine and was quenched by proximity with a complementary guanine. The assay demonstrated that quenching varies linearly with the concentration of template across a defined concentration range. The commonly used fluorophore FITC is inherently quenched by deoxyguanosine nucleotides. The level of quenching can be increased if more guanines are present or a single guanine is located in the first overhang position, 1 nt beyond the fluorophore-labelled terminus of the probe. This approach to amplicon detection is easier to design than fluorogenic oligoprobes, simpler to synthesise and use in real-time PCR and does not require a DNA polymerase with nuclease activity (55). The light-up probe is a peptide nucleic acid to which the asymmetric cyanine fluorophore thiazole orange is attached (56). When hybridised with a nucleic acid target, either as a duplex or triplex, depending on the oligoprobe’s sequence, the fluorophore becomes strongly fluorescent. These probes do not interfere with the PCR, do not require conformational change, are sensitive to single nucleotide mismatches allowing fluorescence melting analysis, and because a single reporter is used, a direct measurement of fluorescence can be made instead of the measurement of a change in fluorescence between two fluorophores (56,57). However, non-specific fluorescence has been reported during later cycles using these probes (58). 5′ Nuclease oligoprobes In the late 1980s homogeneous assays were few and far between, but rapid advances in thermocycler instrumentation and the chemistry of nucleic acid manipulation have since made these assays commonplace. The success of these assays revolves around a signal changing in some rapid and measurable way upon hybridisation of a probe to its target (59). By using an excess, the time required for hybridisation of an oligoprobe to its target, especially when the amount of that target has been increased by PCR or some other amplifying process, is significantly reduced (41,59). In 1991, Holland et al. (6) described a technique that was to form the foundation for homogeneous PCR using fluorogenic oligoprobes. Amplicon was detected by monitoring the effect of Taq DNA polymerase’s 5′→3′ endonuclease activity on specific oligoprobe/target DNA duplexes. The radiolabelled products were examined using thin layer chromatography and the presence or absence of hydrolysis was used as an indicator of duplex formation. These oligoprobes contained a 3′ phosphate moiety, which blocked their extension by the polymerase, but otherwise had no affect on the amplicon’s yield. The desirable criteria for an oligoprobe label are (i) easy attachment of the label to DNA, (ii) detectability at low concentrations, (iii) detectability using simple instrumentation, (iv) production of an altered signal upon specific hybridisation, (v) biological safety, (vi) stability at elevated temperatures and (vii) an absence of interference with the activity of the polymerase (6,18). An innovative approach used nick-translation PCR in combination with dual-fluorophore labelled oligoprobes (14). In the first truly homogenous assay of its kind, one fluorophore was added to the 5′ terminus and one to the middle of a sequence specific oligonucleotide probe. When in such close proximity, the 5′ reporter fluorophore (6-carboxy-fluoroscein) transferred laser-induced excitation energy by FRET to the 3′  quencher fluorophore (6-carboxy-tetramethyl-rhodamine; TAMRA), which reduced the lifetime of the reporter’s excited state by taking its excess energy and emitting it as a fluorescent signal of its own (Fig. 2A and B). TAMRA emitted the new energy at a wavelength that was monitored but not utilised in the presentation of data. However, when the oligoprobe hybridised to its template, the fluorophores were released due to hydrolysis of the oligoprobe component of the probe/target duplex. Once the labels were separated, the reporter’s emissions were no longer quenched and the instrument monitored the resulting fluorescence. These oligoprobes have been called 5′ nuclease, hydrolysis or TaqMan® oligoprobes (Fig. 3A). Nuclease oligoprobes have design requirements that are applicable to the other linear oligoprobe chemistries, including (i) a length of 20–40 nt, (ii) a GC content of 40–60%, (iii) no runs of a single nucleotide, particularly G, (iv) no repeated sequence motifs, (v) an absence of hybridisation or overlap with the forward or reverse primers and (vi) a TM at least 5°C higher than that of the primers, to ensure the oligoprobe has bound to the template before extension of the primers can occur (60). This technology, however, required the development of a platform to excite and detect fluorescence as well as perform thermal cycling. A charge-coupled device had been described in 1992 for the quantification of conventional reverse transcription (RT)–PCR products (61). In 1993 this approach was combined with a thermal cycler resulting in the first real-time PCR fluorescence excitation and detection platform (29). To date, the ABI Prism® 7700 sequence detection system (Perkin Elmer Corporation/Applied Biosystems, USA) has been the main instrument used for 5′ nuclease oligoprobes. Non-PCR related fluorescence fluctuations have been normalised using a non-participating or ‘passive’ internal reference fluorophore (6-carboxy-N,N,N′,N′-tetramethylrhodamine; ROX). The corrected values, obtained from a ratio of the emission intensity of the reporter signal and ROX, are called RQ+. To further control amplification fluctuations, the fluorescence from a ‘no-template’ control reaction (RQ–) is subtracted from RQ+ resulting in the ΔRQ value that indicates the magnitude of the signal generated for the given PCR (62). The fractional cycle number at which the real-time fluorescence signal mirrors progression of the reaction above the background noise was used as an indicator of successful target amplification (63). This threshold cycle (CT) is defined as the PCR cycle in which the gain in fluorescence generated by the accumulating amplicon exceeds 10 standard deviations of the mean baseline fluorescence, using data taken from cycles 3 to 15 (Fig. 1B) (64). The CT is proportional to the number of target copies present in the sample (17). A recent improvement to the nuclease oligoprobe has resulted in the minor groove binding (MGB) oligoprobes (Fig. 3A, inset). This chemistry replaces the standard TAMRA quencher with an NFQ and incorporates a molecule that stabilises the oligoprobe-target duplex by folding into the minor groove of the dsDNA (65). This allows the use of very short (14 nt) oligoprobes, which are ideal for detecting single nucleotide polymorphisms (SNPs). A related use of dual labelled oligonucleotide sequences has been to provide the signal-generating portion of the DzyNA–PCR system (66). Here, the reporter and quencher are separated after cleavage of the probe by a DNAzyme, which is created during PCR as the complement of an antisense DNAzyme sequence included in the 5′ tail of one of the primers. Upon cleavage, the dual labelled substrate releases the fluorophores and generates a signal in an analogous manner to the 5′ nuclease probe. Hairpin oligoprobes Molecular beacons were the first hairpin oligoprobes and are a variation of the dual-labelled nuclease oligoprobe (Fig. 3B). The hairpin oligoprobe’s fluorogenic labels are called fluorophore and quencher, and they are positioned at the termini of the oligoprobe. The labels are held in close proximity by distal stem regions of homologous base pairing deliberately designed to create a hairpin structure which results in quenching either by FRET or a direct energy transfer by a collisional mechanism due to the intimate proximity of the labels (Fig. 2E and F) (67). In the presence of a complementary sequence, designed to occur within the bounds of the primer binding sites, the oligoprobe will hybridise, shifting into an open configuration. The fluorophore is now spatially removed from the quencher’s influence and fluorescent emissions are monitored during each cycle (68). The occurrence of a mismatch between a hairpin oligoprobe and its target has a greater destabilising effect on the duplex than the introduction of an equivalent mismatch between the target and a linear oligoprobe. This is because the hairpin structure provides a highly stable alternate conformation. Therefore, hairpin oligoprobes have been shown to be more specific than the more common linear oligoprobes making them ideal candidates for detecting SNPs (67). The quencher, 4-(4′-dimethylamino-phenylazo)-benzene (DABCYL), differs from that described for the nuclease oligoprobes because it is an NFQ. The wavelength-shifting hairpin probe is a recent improvement to this chemistry which makes use of a second, harvesting fluorophore. The harvester passes excitation energy acquired from a blue light source and releases it as fluorescent energy in the far-red wavelengths. The energy can then be used by a receptive ‘emitter’ fluorophore that produces light at characteristic wavelengths (Fig. 3B, inset). This offers the potential for improved multiplex real-time PCR and SNP analysis (Fig. 4), using currently available instruments (69). Because the function of these oligoprobes depends upon correct hybridisation of the stem, accurate design is crucial to their function (47). Self-fluorescing amplicon The self-priming amplicon is similar in concept to the hairpin oligoprobe, except that the label becomes irreversibly incorporated into the PCR product (Fig. 3D and E). Two approaches have been described: sunrise primers (now commercially called Amplifluor™ hairpin primers) and scorpion primers (31,70). The sunrise primer consists of a 5′ fluorophore and a DABCYL NFQ. The labels are separated by complementary stretches of sequence that create a stem when the sunrise primer is closed. At the 3′ terminus is a target-specific primer sequence. The sunrise primer’s sequence is intended to be duplicated by the nascent complementary strand and, in this way, the stem is destabilised, the two fluorophores are held ~20 nt (70 Å) apart and the fluorophore is free to emit its excitation energy for monitoring (70). This system could suffer from non-specific fluorescence due to duplication of the sunrise primer sequence during the formation of primer-dimer. The scorpion primer is almost identical in design except for an adjacent hexethylene glycol molecule that blocks duplication of the signalling portion of the scorpion. In addition to the difference in structure, the function of scorpion primers differs slightly in that the 5′ region of the oligonucleotide is designed to hybridise to a complementary region within the amplicon. This hybridisation forces the labels apart disrupting the hairpin and permitting emission in the same way as hairpin probes (31). VIRAL QUANTITATION The majority of diagnostic PCR assays reported to date have been used in a qualitative, or ‘yes/no’ format. The development of real-time PCR has brought true quantitation of target nucleic acids out of the pure research laboratory and into the diagnostic laboratory. Determining the amount of template by PCR can be performed in two ways: as relative quantitation and as absolute quantitation. Relative quantitation describes changes in the amount of a target sequence compared with its level in a related matrix. Absolute quantitation states the exact number of nucleic acid targets present in the sample in relation to a specific unit (71). Generally, relative quantitation provides sufficient information and is simpler to develop. However, when monitoring the progress of an infection, absolute quantitation is useful in order to express the results in units that are common to both scientists and clinicians and across different platforms. Absolute quantitation may also be necessary when there is a lack of sequential specimens to demonstrate changes in virus levels, no suitably standardised reference reagent or when the viral load is used to differentiate active versus persistent infection. A very accurate approach to absolute quantitation by PCR is the use of competitive co-amplification of an internal control nucleic acid of known concentration and a wild-type target nucleic acid of unknown concentration, with the former designed or chosen to amplify with an equal efficiency to the latter (72–76). However, while conventional competitive PCR is relatively inexpensive, real-time PCR is far more convenient, reliable and better suited to quick decision making in a clinical situation (77,78). This is because conventional, quantitative, competitive PCR (qcPCR) requires significant development and optimisation to ensure reproducible performance and a predetermined dynamic range for both the amplification and detection components (79). Although a comparison of absolute standard curves, relative standard curves and CT values produces similar final values (80), the general belief remains that an internal control in combination with replicates of each sample are essential for reliable quantitation by PCR (38,39). Unfortunately, real-time PCR software with the ability to calculate the concentration of an unknown by comparing signals generated by an amplified target and internal control is only beginning to emerge. This issue will hopefully be addressed in upcoming commercial releases (81). Therefore, the next best approach to quantitation by PCR is the use of an external standard curve. This approach relies upon titration of an identically amplified template, in a related sample matrix, within the same experimental run. While the external standard curve is the more commonly described approach, it suffers from uncontrolled and unmonitored inter-tube variations. Because of this omission, such experiments should be described as semi-quantitative. Despite this sub-optimal approach, fluorescence data is generally collected from PCR cycles that span the linear amplification portion of the reaction where the fluorescent signal and the accumulating DNA are proportional. Because the emissions from fluorescent chemistries are temperature dependent, data is generally acquired only once per cycle at the same temperature in order to monitor amplicon yield (45). The CT of the sample at a specific fluorescence value can then be compared with similar data collected from a series of standards by the calculation of a standard curve.The determination of the CT depends upon the sensitivity and ability of the instrument to discriminate specific fluorescence from background noise, the concentration and nature of the fluorescence-generating component and the amount of template initially present. Real-time PCR offers significant improvements to the quantitation of viral load because of its enormous dynamic range that can accommodate at least eight log10 copies of nucleic acid template (33,52,77,82–89). This is made possible because the data are chosen from the linear phase (LP; Fig. 1B) of amplification where conditions are optimal, rather than the end-point where the final amount of amplicon present may have been affected by inhibitors, poorly optimised reaction conditions or saturation by inhibitory PCR by-products and double-stranded amplicon. The result of taking data from the end-point is that there may not be a relationship between the initial template and final amplicon concentrations. Real-time PCR is also an attractive alternative to conventional PCR for the study of viral load because of its low inter-assay and intra-assay variability (77,87,90) and its equivalent or greater analytical sensitivity in comparison with traditional viral culture, or conventional single-round, and nested PCR (77,85,91–96). Real-time PCR has been reported to be at least as sensitive as Southern blot (92). However, these reports could be an over-estimate due to the choice of smaller targets, which amplify more efficiently, or due to the use of different or improved primers for the real-time assays because the use of software to design optimised primers and oligoprobes is more common. When this increased sensitivity and broad dynamic range are combined, it is possible to quantitate template from samples containing a large range of concentrations, as is often the case in patient samples. This avoids the need for dilution of the amplicon prior to conventional detection or repeat of the assay using a diluted sample because the first test result falls outside the limits of the assay. These are problems encountered when using some conventional qcPCR assay kits, which cannot encompass high viral loads whilst maintaining suitable sensitivity (52,97–99). The flexibility of real-time PCR is also demonstrated by its ability to detect one target in the presence of a vast excess of another target in duplexed assays (84). Viral load is also a useful indicator of the extent of active infection, virus–host interactions and the response to antiviral therapy, all of which can play a role in the treatment regimen selected (100,101). Conventional quantitative PCR has already proven the benefits of applying nucleic acid amplification to the monitoring of viral load as a useful marker of disease progression and as a component of studies into the efficacy of antiviral compounds (74,100,102–104). The severity of some diseases has been shown to correlate with the viral load making real-time PCR quantitation useful to study not simply the presence of a virus but the role of viral reactivation or persistence in the progression of disease (78,82,91,105–112). An example of the benefits which real-time PCR has brought to the quantitative detection of human cytomegalovirus (CMV) is seen in patients who are immunosuppressed following solid organ or bone marrow transplantation. Although qualitative detection of CMV DNA by PCR has been used as an indicator for the success of antiviral therapy, quantitative assays are preferred in order to monitor patient’s therapeutic responses. Moreover, since it has been postulated that the monitoring of viral replication over time is a more reliable indicator of a developing viral disease than the determination of absolute viral amounts at a single point of time, several quantitative assays have been established and evaluated to increase diagnostic accuracy. Quantitative competitive assays based on end-point analysis have displayed detection limits of 5 × 101 genome equivalents (ge) per assay and a dynamic range of 5 × 101–5 × 104 ge/assay (113). Hybridisation-based assays covered approximately the same dynamic range of four orders of magnitude with detection limits of 20 ge/assay (114,115). Although these assays possess dynamic ranges that may be sufficient for most clinical applications, they display a high inter- and intra-assay variability, up to 40% (115). In contrast, one of the first published real-time PCR assays for the detection of CMV DNA could be performed in <90 min, spanned a dynamic range of six to seven orders of magnitude with a detection limit of at least 10 ge/assay and an inter-assay and intra-assay variability of <10% and <5%, respectively, using plasma samples from bone marrow transplant patients (116). MULTIPLEX REAL-TIME PCR Multiplexing (using multiple primers to allow amplification of multiple templates within a single reaction) is a useful application of conventional PCR (117). However, its transfer to real-time PCR has confused its traditional terminology. The term multiplex real-time PCR is more commonly used to describe the use of multiple fluorogenic oligoprobes for the discrimination of multiple amplicons. The transfer of this technique has proven problematic because of the limited number of fluorophores available (14) and the common use of a monochromatic energising light source. Although excitation by a single wavelength produces bright emissions from a suitably selected fluorophore, this restricts the number of fluorophores that can be included (69). Recent improvements to the design of the hairpin primers, and hairpin and nuclease oligoprobes as well as novel combinations of fluorophores such as in the bi-probe and light-up probe systems, have promised the ability to discriminate an increasing number of targets. The discovery and application of the non-fluorescent quenchers has liberated some wavelengths that were previously occupied by the emissions from the early quenchers themselves. This breakthrough has permitted the inclusion of a greater number of spectrally discernable oligoprobes per reaction, and highlighted the need for a single non-fluorescent quencher, which can quench a broad range of emission wavelengths (e.g. 400–600 nm). Early real-time PCR systems contained optimised filters to minimise overlap of the emission spectra from the fluorophores. Despite this, the number of fluorophores that could be combined and clearly distinguished was limited when compared with the discriminatory abilities of conventional multiplex PCR. More recent real-time PCR platforms have incorporated either multiple light-emitting diodes to span the entire visible spectrum, or a tungsten light source, which emits light over a broad range of wavelengths. When these platforms incorporate high quality optical filters it is possible to apply any current real-time PCR detection chemistries on the one machine. Nonetheless, these improvements generally allow only four-colour oligoprobe multiplexing, of which one colour is ideally set aside for an internal control to monitor inhibition and perhaps even act as a co-amplified competitor. Some real-time PCR designs have made use of single or multiple nucleotide changes between similar templates to allow their differentiation by TM (Fig. 4) thus avoiding the need for multiple fluorophores (49,91,118–122). This approach has been used far more commonly in the detection of human genetic diseases where as many as 27 possible nucleotide substitutions have been detected using only one or two fluorophores (48,123–128). To date, there have only been a handful of truly multiplexed real-time PCR assays described in the literature and few of these have been applied to the diagnosis of infectious disease. Some of these approaches cannot, technically, be considered real-time, homogenous assays because they require interruption of the procedure to transfer template, their fluorescence is detected by end-point analysis or the assays are not performed within the same tube. One of the best viral, multiplex, real-time PCR protocols can discriminate between four retroviral target sequences (129), however, conventional multiplex PCR using end-point detection can easily discriminate more than five different amplified sequences, indicating a greater flexibility when compared with real-time PCR (130–133). Despite these limitations, a number of assays have been applied to the detection of several viral genomes at once, including the use of non-specific label, SYBR green 1 to detect herpes simplex viruses (HSV), varicella zoster virus (VZV) or CMV, in separate tubes (134) or, by adaptation of a conventional multiplex PCR, to identify HSV-1 and HSV-2, VZV and enteroviruses within a single capillary by applying fluorescent melting curve analysis (121). Another single-tube multiplexed nuclease oligoprobe RT–PCR was capable of simultaneously detecting Influenza A and B in patient’s respiratory samples with improved sensitivity compared with conventional or shell-viral culture (95). Future developments of novel chemistries such as combinatorial fluorescence energy transfer tags (135), and improvements to the design of real-time instrumentation and software will greatly enhance the future of multiplex real-time PCR. APPLICATIONS TO VIROLOGY Real-time PCR has been extremely useful for studying viral agents of infectious disease and helping to clarify disputed infectious disease processes. Most of the assays presented in the literature allow an increased frequency of virus detection compared with conventional techniques, which makes the implementation of real-time PCR attractive to many areas of virology. Of course, real-time PCR has proven increasingly valuable for general virological studies, although increasingly, these applications are difficult to review due to the nature of their use as a tool rather than the focus of published study. Such studies have investigated the role of viruses in a range of diseases by simply confirming the presence or absence of the virus (136,137) or, in the future, by monitoring the levels of specific gene activity (138) as a result of growth under manipulated conditions. Altered viral entry or replication, caused by the modification of target tissues, can also be followed using real-time PCR as can links between virus replication and the expression of cellular genes (139–141). Real-time PCR has enhanced the speed and scope of measuring viral strain and titre differences in patients displaying different syndromes due to varieties of the same virus (106). Also, epidemiological studies have been improved in speed and scope through the use of real-time PCR because it can reliably measure the amount of two nucleic acid targets within a single reaction (91,142). New chemistries have allowed better discrimination of multiple viral genotypes within a single reaction vessel (143) and provided an alternative to methods of virus detection based on morbidity and mortality assays. The use of real-time PCR has provided insight into the role(s) some compounds have on PCR inhibition as well as shedding light on the efficiency of different nucleic acid extraction methods, from a diverse range of sample types (144–146). This ability to utilise template from a range of sample types fulfils a requirement for an ideal detection system, which is the ability to apply a single technology across many fields. This flexibility is highlighted by the detection of viral nucleic acids derived, in different ways, from plants (147–149), animals (86,89,94,150–152), urban sludge (85,153), tissue culture (23,77,96,154–160), various solid tissues (108,118,161–166), cerebrospinal fluid (49,167,168), peripheral blood mononuclear cells (82,93,105,112,169–171), plasma (81,88,90,172–176), serum (30,33,36,87,97,99,109,177–181), swabs (182,183), saliva (106) and urine (78,122,184,185). Also, chronic conditions such as sarcoma (186–189), carcinoma (92,111), cervical intraepithelial neoplasia (190–192), and lymphoproliferative disorders (193,194) can be relatively easily studied to investigate direct or indirect links with viral infection. These studies have targeted viruses which include the flaviviruses (33,36,96,109,195), hepadnaviridae (52,97), herpesviruses (21,77,78,82–84,91,92,98,105,106,108, 111,112,144,167,171), orthomyxoviruses (95), parvoviruses (88), papovaviruses (122,143,145), paramyxoviruses (94), picornaviruses (85,86), retroviruses (90,93,156,196,197) and TT virus (137). Viral load monitoring by real-time PCR has also proven beneficial for studying patients following organ transplant (21,83,107,198–201). This technology has now become an essential tool in the thorough assessment of viral gene therapy vectors prior to their use in clinical trials. Nuclease oligoprobes have been most commonly reported for these studies, which assess the biodistribution, function and purity of these drug preparations (164,197,202–205). Additionally, the study of emerging viruses has been complimented by the use of real-time PCR as a tool to demonstrate links between unique viral sequences and patient clinical signs and symptoms (94,96,150,160,206,207). The speed and flexibility of real-time PCR has also proven useful to commercial interests for the screening of microbial contamination of large-scale reagent preparations produced from eukaryotic expression systems (208,209). CONCLUSIONS AND SUMMARY Advances in the development of fluorophores, nucleotide labelling chemistries, and the novel applications of oligoprobe hybridisation have provided real-time PCR technology with a broad enough base to ensure its acceptance. Recently, instrumentation has appeared that is capable of incredibly short cycling times combined with the ability to detect and differentiate multiple amplicons. New instruments are also flexible enough to allow the use of any of the chemistries described in this review making real-time nucleic acid amplification an increasingly attractive and viable proposition for the routine diagnostic laboratory. In many cases these laboratories perform tissue culture to isolate virus and serological methods to confirm the identity of the isolate, which may take a considerable, and clinically relevant, amount of time. The familiarity that leads to comfortable routine use of a technology is now apparent in the inclusion of the fluorogenic oligoprobe chemistries in many laboratories. According to the literature, the most widely used format is the 5′ endonuclease oligoprobe although that is most likely due to its commercial maturity. The more recently developed oligoprobe chemistries have been used in a number of innovative applications and it is apparent from the rate and content of real-time PCR-related publications that they are becoming more widely accepted. Recent developments in multiplex real-time PCR have suggested a future in which easy identification, genotyping and quantitation of viral targets in single, rapid reactions will be commonplace. Of course, this technology is by no means restricted to virology, as significant achievements have appeared in the area of mutation detection, applying all the benefits described above to enhance the detection of genetic disease and, where applicable, allow quantitation of the extent of such genetic changes. Micro- and macro-array technology will surely play some chimeric role in the real-time PCR of future research, but for now there is significant potential for routine, research and commercial interests to redesign existing systems for greater sophistication, flexibility and the ability to generate high quality quantitative data. The development of assays capable of real-time PCR that can discriminate as many targets as conventional multiplex PCR assays, whilst producing quantitative data at a greatly increased speed, will consolidate fluorogenic nucleic acid amplification as a routine tool for the laboratory of tomorrow. ACKNOWLEDGEMENTS The Royal Children’s Hospital Foundation Grant number 922-034, which was sponsored by the Woolworth’s ‘Care for Kids’ campaign, supported this work. This is publication number 139 from the Sir Albert Sakzewski Virus Research Centre. * To whom correspondence should be addressed at: CVRU, SASVRC, Royal Children’s Hospital, Herston Road, Herston, Queensland 4029, Australia. Tel: +61 3636 8716; Fax: +61 3636 1401; Email: [email protected] View largeDownload slide Figure 1. Kinetic amplification. (A) An idealised plot of temperature versus time during a single PCR cycle comprised of the denaturation (D), primer and probe annealing (A) and primer extension (E) steps. At the indicated optimal temperature ranges, dsDNA is denatured (TD), oligoprobes anneal (TM-PROBE) and finally the primers anneal as a precursor to their extension (TM-PRIMER). The actual temperature, shown as a dashed line, may overshoot the desired temperature to varying degrees, depending on the quality of the thermocycler employed. (B) The ideal amplification curve of a real-time PCR (bold), when plotted as fluorescence intensity against the cycle number, is a typical sigmoidal growth curve. Early amplification cannot be viewed because the detection signal is indistinguishable from the background. However, when enough amplicon is present, the assay’s exponential progress can be monitored as the rate of amplification enters a log-linear phase (LP). Under ideal conditions, the amount of amplicon increases at a rate of approximately one log10 every three cycles. As primers and enzyme become limiting and products inhibitory to the PCR accumulate, the reaction slows, entering a transition phase (TP), eventually reaching the plateau phase (PP) where there is little or no further increase in product yield. The point at which the fluorescence passes from insignificant levels to clearly detectable is called the threshold cycle (CT; indicated by an arrow), and this value is used in the calculation of template quantity during quantitative real-time PCR. Also shown are curves representing an optimal titration of template (grey), consisting of higher and lower starting template concentrations, which produce lower or higher CT values, respectively. Data for the construction of a standard curve are taken from the LP, which subsequently allows the concentration of unknown samples to be determined. View largeDownload slide Figure 1. Kinetic amplification. (A) An idealised plot of temperature versus time during a single PCR cycle comprised of the denaturation (D), primer and probe annealing (A) and primer extension (E) steps. At the indicated optimal temperature ranges, dsDNA is denatured (TD), oligoprobes anneal (TM-PROBE) and finally the primers anneal as a precursor to their extension (TM-PRIMER). The actual temperature, shown as a dashed line, may overshoot the desired temperature to varying degrees, depending on the quality of the thermocycler employed. (B) The ideal amplification curve of a real-time PCR (bold), when plotted as fluorescence intensity against the cycle number, is a typical sigmoidal growth curve. Early amplification cannot be viewed because the detection signal is indistinguishable from the background. However, when enough amplicon is present, the assay’s exponential progress can be monitored as the rate of amplification enters a log-linear phase (LP). Under ideal conditions, the amount of amplicon increases at a rate of approximately one log10 every three cycles. As primers and enzyme become limiting and products inhibitory to the PCR accumulate, the reaction slows, entering a transition phase (TP), eventually reaching the plateau phase (PP) where there is little or no further increase in product yield. The point at which the fluorescence passes from insignificant levels to clearly detectable is called the threshold cycle (CT; indicated by an arrow), and this value is used in the calculation of template quantity during quantitative real-time PCR. Also shown are curves representing an optimal titration of template (grey), consisting of higher and lower starting template concentrations, which produce lower or higher CT values, respectively. Data for the construction of a standard curve are taken from the LP, which subsequently allows the concentration of unknown samples to be determined. View largeDownload slide Figure 2. Fluorogenic mechanisms. When a 5′ nuclease probe’s reporter (R) and quencher (Q, open) are in close proximity as in (A), the quencher ‘hijacks’ the emissions that have resulted from excitation of the reporter by the instrument’s light source. The quencher then emits this energy (solid arrows). When the fluorophores are separated beyond a certain distance, as occurs upon hydrolysis as depicted in (B), the quencher no longer exerts any influence and the reporter emits at a distinctive wavelength (dashed arrows) which is recorded by the instrument. In the reverse process as depicted in (C) using adjacent oligoprobes, the fluorophores begin as separated entities. A signal from the acceptor (A) can only be generated when the donor (D) comes into close proximity as shown in (D). This occurs as the result of adjacent hybridisation of the oligoprobes to the target amplicon. In (E) another form of quenching is shown, caused by the intimate contact of labels attached to hairpin oligoprobes (molecular beacon, sunrise or scorpion). The fluorophore (F) and a NFQ (Q, closed) interact more by collision than FRET, disrupting each other’s electronic structure and directly passing on the excitation energy which is dissipated as heat (jagged, dashed arrows). When the labels are separated by disruption of the hairpin, as is the case in (F), the fluorophore is free to fluoresce (dashed arrows). View largeDownload slide Figure 2. Fluorogenic mechanisms. When a 5′ nuclease probe’s reporter (R) and quencher (Q, open) are in close proximity as in (A), the quencher ‘hijacks’ the emissions that have resulted from excitation of the reporter by the instrument’s light source. The quencher then emits this energy (solid arrows). When the fluorophores are separated beyond a certain distance, as occurs upon hydrolysis as depicted in (B), the quencher no longer exerts any influence and the reporter emits at a distinctive wavelength (dashed arrows) which is recorded by the instrument. In the reverse process as depicted in (C) using adjacent oligoprobes, the fluorophores begin as separated entities. A signal from the acceptor (A) can only be generated when the donor (D) comes into close proximity as shown in (D). This occurs as the result of adjacent hybridisation of the oligoprobes to the target amplicon. In (E) another form of quenching is shown, caused by the intimate contact of labels attached to hairpin oligoprobes (molecular beacon, sunrise or scorpion). The fluorophore (F) and a NFQ (Q, closed) interact more by collision than FRET, disrupting each other’s electronic structure and directly passing on the excitation energy which is dissipated as heat (jagged, dashed arrows). When the labels are separated by disruption of the hairpin, as is the case in (F), the fluorophore is free to fluoresce (dashed arrows). View largeDownload slide Figure 3. Oligoprobe chemistries. (A) 5′ Nuclease oligoprobes. As the DNA polymerase (pol) progresses along the relevant strand, it displaces and then hydrolyses the oligoprobe via its 5′→3′ endonuclease activity. Once the reporter (R) is removed from the extinguishing influence of the quencher (Q, open), it is able to release excitation energy at a wavelength that is monitored by the instrument and different from the emissions of the quencher. Inset shows the NFQ and MGB molecule that make up the improved MGB nuclease-oligoprobes. (B) Hairpin oligoprobes. Hybridisation of the oligoprobe to the target separates the fluorophore (F) and non-fluorescent quencher (Q, closed) sufficiently to allow emission from the excited fluorophore, which is monitored. Inset shows a wavelength-shifting hairpin oligoprobe incorporating a harvester molecule. (C) Adjacent oligoprobes. Adjacent hybridisation results in a FRET signal due to interaction between the donor (D) and acceptor (A) fluorophores. This bimolecular system acquires its data from the acceptor’s emissions in an opposite manner to the function of nuclease oligoprobe chemistry. (D) Sunrise primers. The opposite strand is duplicated so that the primer’s hairpin structure can be disrupted. This separates the labels, eliminating the quenching in a similar manner to the hairpin oligoprobe. (E) Scorpion primers. The primer does not require extension of the complementary strand; in fact it blocks extension to ensure that the hairpin in the probe is only disrupted by specific hybridisation with a complementary sequence designed to occur downstream of its own, nascent strand. Inset shows a duplex scorpion that exchanges the stem–loop structure for a primer element terminally labelled with the fluorophore and a separate complementary oligonucleotide labelled with a quencher at the 5′ terminus. View largeDownload slide Figure 3. Oligoprobe chemistries. (A) 5′ Nuclease oligoprobes. As the DNA polymerase (pol) progresses along the relevant strand, it displaces and then hydrolyses the oligoprobe via its 5′→3′ endonuclease activity. Once the reporter (R) is removed from the extinguishing influence of the quencher (Q, open), it is able to release excitation energy at a wavelength that is monitored by the instrument and different from the emissions of the quencher. Inset shows the NFQ and MGB molecule that make up the improved MGB nuclease-oligoprobes. (B) Hairpin oligoprobes. Hybridisation of the oligoprobe to the target separates the fluorophore (F) and non-fluorescent quencher (Q, closed) sufficiently to allow emission from the excited fluorophore, which is monitored. Inset shows a wavelength-shifting hairpin oligoprobe incorporating a harvester molecule. (C) Adjacent oligoprobes. Adjacent hybridisation results in a FRET signal due to interaction between the donor (D) and acceptor (A) fluorophores. This bimolecular system acquires its data from the acceptor’s emissions in an opposite manner to the function of nuclease oligoprobe chemistry. (D) Sunrise primers. The opposite strand is duplicated so that the primer’s hairpin structure can be disrupted. This separates the labels, eliminating the quenching in a similar manner to the hairpin oligoprobe. (E) Scorpion primers. The primer does not require extension of the complementary strand; in fact it blocks extension to ensure that the hairpin in the probe is only disrupted by specific hybridisation with a complementary sequence designed to occur downstream of its own, nascent strand. Inset shows a duplex scorpion that exchanges the stem–loop structure for a primer element terminally labelled with the fluorophore and a separate complementary oligonucleotide labelled with a quencher at the 5′ terminus. View largeDownload slide Figure 4. Fluorescent melting curve analysis. At the completion of a real-time PCR using a pair of adjacent oligoprobes, the reaction can be cooled to a temperature below the expected TM of the oligoprobes then heated above 85°C at a fraction of a degree per second (B). During heating, the emissions of the acceptor can be constantly acquired (C). Software calculates the negative derivative of the fluorescence over time, producing clear peaks that indicate the TM of the oligoprobe-target melting transition (D). When one or more mutations are present under one or both oligoprobes (A), this TM is shifted and this shift can be used diagnostically to discriminate single nucleotide polymorphisms in the template. Ideally, one of the oligoprobes, the anchor, is designed to bind to a stable sequence region, whereas the other, sensor, will span the mismatch. Mismatches near the centre of the probe and flanked by G:C pairs are more destabilising than mismatches near the ends of the oligoprobe flanked by A:T pairs. View largeDownload slide Figure 4. Fluorescent melting curve analysis. At the completion of a real-time PCR using a pair of adjacent oligoprobes, the reaction can be cooled to a temperature below the expected TM of the oligoprobes then heated above 85°C at a fraction of a degree per second (B). 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Badi, Laura;Barberis, Alcide

doi: 10.1093/nar/30.6.1306pmid: 11884627

Abstract Activation of transcription in eukaryotes requires the concerted action of numerous components of the RNA polymerase II transcriptional apparatus. The degree of dependence on many of these components varies from gene to gene and it is still largely unknown how the requirement for any particular component is determined at any given gene. We show that removal of Gal11 from the yeast transcription complex can affect activation from the CUP1 UAS in a manner dependent on its genomic context. Our results indicate a novel function for the CUP1 upstream repeated element (CURE) located upstream of the CUP1 UAS at the naturally multimerized CUP1 locus. The presence of CURE endowed the CUP1 UAS with a reduced susceptibility to the effects of deleting Gal11. Similar results were obtained with the Srb/mediator subunit Srb5. Restoration of activation from the CUP1 promoter to wild-type levels by the CURE correlated with changes in the accessibility of local chromatin to nucleases. The CURE sequence may serve to protect the stress-inducible CUP1 UAS–promoter elements against reduced activation that may result from crippled transcription complexes under stress conditions. Received December 22, 2001; Accepted January 21, 2002. INTRODUCTION Genes transcribed by RNA polymerase II (pol II) are generally silent unless activated by sequence-specific DNA-binding proteins called transcriptional activators. The function of activators at specific promoters can be counteracted by another class of DNA-binding proteins known as gene-specific repressors. Despite the pivotal role of these two types of regulatory proteins in controlling the process of differential gene expression (1,2), it is believed that they alone cannot account for the high degree of specificity and coordination of gene regulation in any given eukaryotic cell at any time during the cell cycle or in rapid response to changes in the environment (3–5). The eukaryotic transcriptional machinery is composed of a large number of proteins that include, in addition to the pol II core enzyme, the general transcription factors, the Srb/mediator complex, the Srb10 CDK complex, the SAGA complex and the Swi/Snf complex, among others (3). It is conceivable that the raison d’être of such a large and complicated apparatus in eukaryotes (as compared to the relatively simple transcriptional apparatus in bacteria) is linked to the complexity of the combinatorial control of gene expression in eukaryotic cells (4). Recent results have shown that the RNA pol II transcription complex participates in transcriptional regulation by providing different activities that can be targets of certain signaling pathways and ultimately modulate the action of the pol II core enzyme (3). Genome-wide analysis of gene expression in yeast cells carrying mutations in a variety of components of the pol II transcription complex has shown that the requirement for many components varies from gene to gene (5). For example, while Srb4 protein is required for full expression of ∼93% of yeast genes, Srb5, a partner of Srb4 in the Srb/mediator complex, is only required at ∼16% of genes. In addition to simple requirement, the role of some of these components in the expression of a subset of genes may also vary. For example, while the chromatin remodeling Swi/Snf complex is required for activation of ∼2% of yeast genes, other genes accounting for ∼4% of yeast genes appear to be negatively regulated by this complex and thus show elevated expression in its absence (5). An additional example of a factor with a potential dual function is provided by Gal11, a component of the Srb/mediator complex. Gal11 is needed for full expression of ∼40% of yeast genes, while its deletion causes an increase in the expression of ∼3% of yeast genes (6). It is still largely unknown how the requirement for any particular component and its role is determined at any given gene. In this work we have tested the possible role of chromosomal context in determining the dependence on the transcription factor Gal11 for activation of transcription from the CUP1 promoter. CUP1 was chosen because activation of transcription from this promoter seems to be exceptionally unaffected by the lack of transcription factors that are otherwise required for the activation of most yeast genes. For example, conditional ablation of the essential and widely required Srb4 protein and the TFIIH kinase subunit Kin28 or partial deletion of the C‐terminal domain of the pol II enzyme did not affect transcription from the CUP1 promoter (7–9). The CUP1 gene encodes a metallothionein protein and is induced by stress conditions such as high levels of copper (Cu2+) and heat shock (10,11). Inducible and rapid production of the Cup1 protein in yeast is achieved by a combination of two systems: amplification of CUP1 gene copy number in the genome and its transcriptional regulation. The CUP1 locus contains a cluster of tandemly repeated units, which can be found in as many as 15 linearly arrayed copies in some yeast strains (12). Transcription from the CUP1 promoter is activated by sequence-specific transcription factors that bind the CUP1 upstream activating sequence (CUP1 UAS) (13). This sequence contains at least three binding sites for the Cu2+-dependent transcriptional activator Ace1/Cup2 (10) and one binding site for the heat shock-responsive factor Hsf1 (14). The CUP1 UAS is necessary and sufficient for rapid and robust activation of transcription (10,14). The results presented here show that removal of Gal11 from the transcription complex significantly affected activation of transcription from the CUP1 UAS in a manner that was dependent on the genomic context surrounding the reporter gene. Further analysis of these genomic regions revealed a novel role for a DNA sequence in modulating CUP1 activation. This sequence, which we named the CUP1 upstream repeated element (CURE), is naturally present upstream of the CUP1 UAS and is co-amplified with the entire CUP1 transcription unit at the CUP1 locus (see Fig. 3B). We have found that the presence of CURE restored the otherwise impaired activation of transcription by the downstream CUP1 UAS in strains lacking Gal11. This effect was not specific for a Gal11 deletion, since the CURE also restored activation from the CUP1 promoter in the absence of Srb5. We suggest that this novel regulatory element may ensure efficient CUP1 transcription when cells experience stress conditions that may impair the function of some transcription factors. MATERIALS AND METHODS Strains and genetic techniques Yeast genetic techniques and media were as described (15). Yeast transformation was performed by the lithium acetate procedure (16). The Saccharomyces cerevisiae strains used in this study are shown in Table 1. Mutant strains JPY8 (gal11Δ) (17) and BLY30 (srb5Δ) (this study) were obtained from the wild-type strain JPY5 (17) through replacement of the chromosomal GAL11 or SRB5 coding sequences, respectively, with a TRP1 targeting cassette. All reporter strains listed in Table 1 were originated through transformation of the strain of interest with the corresponding integrative reporter plasmids and subsequent selection on leu– selective plates. For each of the integrative plasmids described in Table 2XbaI linearization allowed insertion of the reporters at the CUP1 locus, while BstEII digestion made the reporters suitable for insertion at the LEU2 locus. After transformation with the targeting construct and growth on selective plates, the correct identity of each strain was confirmed by PCR and Southern blot analysis of purified genomic DNA (18). Plasmids Plasmids pBL11 and pBL12 are based on YCplac33 and contain the genome-derived expression cassettes for the GAL11 and SRB5 genes, respectively (19). A list of the episomal and integrative reporter constructs used in this study is shown in Table 2. These plasmids are based on the LEU2-marked centromeric vector pSAL1::lacZ (a kind gift of A. Covarrubias, Institute of Biotechnology, UNAM, Cuernavaca), which contains the Cu2+-inducible promoter region (CUP1 UAS and core promoter) from –394 to +37 bp relative to the first (+1) of the two major transcription start sites of the CUP1 gene fused to the lacZ ORF (20). pSAL1::lacZint is an integrating derivative of pSAL1::lacZ obtained after removal of the ARS/CEN sequence. Plasmid pSAL1–1441::lacZint contains the CUP1 upstream repeated element (–1441 to –395) inserted at its natural location upstream of the CUP1 UAS–promoter region of the lacZ reporter gene. The sequences of all pSAL1::lacZ derivatives were checked for correct reconstitution of the CUP1 regulatory regions. Detailed plasmid maps are available upon request. RNA analysis Yeast total RNA was extracted from exponentially growing cells exposed for 1 h to 400 µm CuSO4 as inducing agent for the CUP1 promoter. For northern blot analysis (21), equal amounts (2.5 µg) of total RNA were assayed to estimate the quantity of specific transcripts derived from the lacZ reporter and from the SNR6 endogenous gene, which is constitutively transcribed by RNA polymerase III. The 32P-labeled probes used in these experiments were the full-length lacZ, CUP1 or SNR6 genes. S1 mapping was performed as described (22) using the lacZ-specific 32P-end-labeled oligonucleotide 5′-cgcacttttcggccaatggtcttggtaattcctttgcgctagaattgaactcaggtacaatcacttcttctgaatgagatttagtcatatatcaa. Quantitation of the radioactive signals was performed with the ImageQuant package (Molecular Dynamics). In all types of RNA mapping experiments the quantity of total input RNA, which was determined by measuring absorption at 260 nm and by ethidium bromide staining of a gel upon electrophoresis, was kept constant and used to normalize the specific signal. β-Galactosidase liquid culture assay The β-galactosidase liquid assay was performed as previously described (23). Yeast cells were exposed to 400 µm CuSO4 for 2 h to induce lacZ expression from the CUP1 promoter before protein extraction. Micrococcal nuclease and restriction endonuclease analysis of intact chromatin Nuclei from exponentially growing yeast cell cultures were extracted as described (24). For micrococcal nuclease (MNase) analysis, nuclei were digested with MNase (4 and 7 U/reaction) for 5 min. Deproteinized DNA samples (naked DNA) were processed in parallel as a control. Nuclei derived from 50 mg of cells (wet weight) were processed in a restriction endonuclease digestion by incubation for 2 h with 10 U of the chosen restriction enzyme. In both experiments, the DNA was purified and used in a secondary digestion reaction with ScaI/PvuII. The fragments of interest were identified through Southern blotting using a 32P-labeled probe obtained by PCR amplification of the NotI–PvuII fragment mapping within the lacZ gene of pSAL1::lacZ (indirect end-labeling technique). RESULTS The negative effect of a GAL11 deletion on CUP1 promoter activation depends on the chromosomal context of the UAS within the CUP1 repeat cluster Biochemical analysis of the Srb/mediator complex has indicated that Gal11 is part of the Gal11/Rgr1/Sin4 subcomplex. Proteins of this subcomplex, unlike other Srb/mediator complex components, have been reported to be relevant cofactors for CUP1 activation (8). Our original interest was to investigate the mechanisms of transcriptional activation from the CUP1 promoter in gal11Δ yeast cells. Measurement and comparison of the CUP1 mRNA levels from Gal11– and Gal11+ cells revealed that the absence of this transcription factor did not significantly affect expression of the endogenous CUP1 genes (Fig. 1A). To restrict the analysis to promoter function, we chose to quantitate the effects of the gal11Δ mutation on transcription levels from a series of lacZ reporter gene constructs based on the CUP1 UAS–promoter elements. The use of such a reporter system also allowed us to distinguish between transcripts produced upon activation of the analyzed CUP1 UAS from those synthesized from the highly duplicated endogenous CUP1 locus. The yeast strains JPY5 (wt) and JPY8 (gal11Δ), which are isogenic except for the GAL11 locus, were transformed with the episomal reporter construct pSAL1::lacZ (see Materials and Methods). Expression of this lacZ reporter gene is controlled by a span of DNA that contains the Cu2+-inducible CUP1 UAS and promoter sequences from –394 to +37 bp relative to the first (+1) of the two major CUP1 transcription start sites (13,25). Measurement of β-galactosidase activity in protein extracts from these transformed strains revealed that the gal11Δ mutant had severely impaired Cu2+-induced lacZ expression levels in comparison to the wild-type strain. The extent of reduction in β-galactosidase activity caused by the lack of Gal11 was ∼80%, as shown in Figure 1B. This decrease in β-galactosidase activity was the direct consequence of a diminished level of Cu2+-induced transcription from the CUP1 UAS::lacZ reporter gene, as shown by measuring the relative lacZ mRNA levels by northern blot assay (Fig. 1C). We therefore conclude that Gal11 is necessary for efficient transcription of a CUP1 UAS::lacZ reporter gene located on an episomal plasmid. We next determined whether Gal11 was also required when the CUP1 UAS::lacZ reporter construct described above was integrated at the natural CUP1 locus within the chromosome. In order to target integration of this reporter gene to the CUP1 chromosomal region, we transformed JPY8 cells (gal11Δ) with an integrative plasmid (pSAL1::lacZint) that had been linearized by digestion at the single XbaI site in the CUP1 UAS sequence (see Materials and Methods). Homologous recombination was expected to occur between the XbaI-digested construct and any one, or more than one, of the repeated CUP1 genes in the yeast genome. Nine yeast clones that had undergone integration of pSAL1::lacZint were selected and analyzed for a requirement for Gal11 for induced activation of the various integrated CUP1 UAS::lacZ reporter genes. When we compared the Cu2+-induced β-galactosidase activities on a plate by colorimetric assay in strains containing either plasmid pBL11 (expressing Gal11) or YCplac33 (the empty parental plasmid) we observed that full activation of the lacZ reporter gene significantly required the presence of Gal11 in two of the nine yeast strains, while relative expression of the reporter gene in seven strains did not seem to be diminished in the absence of Gal11 (data not shown). Moreover, the Gal11-sensitive strains showed a stronger maximal level of lacZ expression than the Gal11-insensitive strains. Figure 2A presents the results of the β-galactosidase assays performed with strains BLY91 (one of the Gal11-sensitive reporter strains) and BLY97 (one of the Gal11-insensitive reporter strains). These results show that the absence of Gal11 caused a 3-fold reduction in Cu2+-induced CUP1 UAS::lacZ reporter gene expression in the BLY91 strain, while it did not significantly affect expression of the reporter gene in strain BLY97. We also compared the levels of Cu2+-induced transcription in these two strains by performing S1 mapping analysis of the lacZ transcript (Fig. 2B). The results of these experiments show that the variations in lacZ mRNA levels upon copper induction correlated with the changes in β-galactosidase activity described above. S1 mapping analysis of the lacZ transcript under non-inducing conditions showed that the CUP1 UAS::lacZ basal expression level in the two strains followed a similar pattern of Gal11 dependence as the induced level of transcription (Fig. 2B). The results of the S1 mapping assays were confirmed by primer extension analysis of the lacZ mRNA (data not shown). The data from the S1 mapping and primer extension analyses of Cu2+-induced UAS::lacZ gene transcription were combined and the average reduction in transcription in gal11Δ cells was calculated as a percentage of expressed lacZ in Gal11– relative to Gal11+ cells. These values were 35.1 ± 0.3% for BLY91 and 87.4 ± 6.3% for BLY97. The CUP1 UAS::lacZ reporter genes in strains BLY91 and BLY97 showed remarkable differences both in their dependence on Gal11 for maximal expression and in the absolute levels of lacZ transcripts that were synthesized. One possible cause for such differences might be the precise location within the CUP1 repeat cluster of the integrated reporter plasmids and the potentially variable copy number. Indeed, the highly duplicated CUP1 locus provides several potential sites for integration of the reporter construct. We therefore performed Southern blot analysis of reporter constructs integrated at the CUP1 locus of strains BLY91 and BLY97. Genomic DNA was prepared from both strains and double digested with KpnI and ScaI restriction nucleases. Upon electrophoresis and blotting, the resultant filters were probed with a radiolabeled CUP1 UAS fragment (see Materials and Methods). Figure 3A shows that the two strains differed in the pattern of integration of the pSAL1::lacZint reporter construct within the CUP1 locus. The deduced pattern of integration is outlined in Figure 3B. Both the BLY91 and BLY97 strains had integrated the CUP1 UAS::lacZ reporter construct at the CUP1 locus, although the two strains displayed differences in copy number and chromosomal context of the integrated plasmid. The BLY91 strain had integrated at least two copies of the lacZ reporter construct that generated a cluster of direct repeats, while the BLY97 strain had inserted a single copy of the lacZ reporter at the 3′-most position available for homologous recombination within the highly duplicated CUP1 locus. This integration site could be identified as the CUP1 UAS belonging to the most downstream CUP1 transcription unit of the repeat (Fig. 3B). Southern blot analysis of the other CUP1 reporter strains revealed that the additional Gal11-sensitive strain, similarly to BLY91, had also integrated at least two copies of the lacZ reporter construct as direct repeats, while the remaining six Gal11-insensitive strains had integrated single copies of the reporter plasmid in one of the endogenous CUP1 genes (data not shown). These results, taken together, suggest that the organization of the reporter plasmid within the CUP1 locus and its copy number might influence the dependence on Gal11 of expression of the CUP1 UAS::lacZ gene. The CUP1 upstream repeated element drastically reduces the dependence on Gal11 for transcriptional activation from the CUP1 UAS The comparative analysis of the CUP1 UAS::lacZ chromosomal locations described above suggested that the different DNA regions present upstream of the respective CUP1 UAS::lacZ genes might determine the difference between these Gal11-sensitive and Gal11-insensitive reporter strains. Since the Gal11-sensitive BLY91 strain contains at least two copies of the pSAL1::lacZint construct in consecutive positions (Fig. 3B), one or more copies of the reporter CUP1 UAS elements are located immediately downstream of plasmid DNA sequences. These CUP1 UAS elements are therefore located in the same sequence context as the CUP1 UAS of the episomal reporter construct described above (see Fig. 1B). In contrast, the CUP1 UAS of the single pSAL1::lacZint copy in the Gal11-insensitive BLY97 strain is located immediately downstream of the chromosomal DNA sequence that is naturally present upstream of the endogenous CUP1 genes. As indicated in Figure 3B, these sequences correspond to the ORF denoted YHR054C, which is part of the CUP1 amplification unit, and which we refer to as the CURE. Thus, a plausible hypothesis is that the CURE sequence is necessary and sufficient to counteract the sensitivity of the CUP1 UAS to deletions in GAL11 (gal11Δ). To test this hypothesis, we reconstituted the different reporter gene structures observed in BLY91 and BLY97 elsewhere in the yeast genome. We engineered a new integrative reporter plasmid, pSAL1–1441::lacZint, by inserting the CURE sequence upstream of the CUP1 UAS in the original pSAL1::lacZint plasmid. This cloning procedure generated the same sequence as found up to 1441 bp upstream of the transcription initiation site of the BLY97 reporter gene. Both plasmids were linearized by digestion at a unique BstEII site within the LEU2 marker gene of the plasmid to target homologous integration at the LEU2 locus of the JPY8 (gal11Δ) strain (see Materials and Methods). The resulting strains were named BLY18, which carries the original pSAL1::lacZint reporter gene integrated at the LEU2 locus, and BLY22, which carries the pSAL1–1441::lacZint reporter construct (including the CURE sequence) at the same chromosomal site. We then measured the transcriptional activity of Cu2+-activated lacZ reporter genes in BLY18 and BLY22 cells transformed with either a Gal11-expressing or an empty plasmid. Reporter gene transcription was monitored by measuring β-galactosidase activity (Fig. 4A) and by northern blot analysis (Fig. 4B). This analysis revealed that full activation of the CUP1 UAS::lacZ reporter gene that lacks the CURE sequence was strongly dependent on Gal11 in strain BLY18. In contrast, in the BLY22 strain the presence of the CURE sequence upstream of the CUP1 UAS element almost completely abolished the dependence on Gal11 for full transcriptional activation (Fig. 4A and B). The CURE drastically reduced the Gal11 requirement without significantly affecting the maximal level of reporter gene expression observed in the presence of Gal11 (Fig. 4A and B). The results of these experiments show that the CURE sequence, which is naturally located upstream of the CUP1 UAS, renders activation of transcription from this promoter almost completely insensitive to the lack of Gal11 in a manner that is independent of the surrounding chromosomal context of the CUP1 locus. The CURE sequence renders activation of transcription from the CUP1 UAS–promoter region independent of Srb5 To follow up the work on Gal11, we tested whether the presence of the CURE sequence upstream of the CUP1 UAS could rescue a transcriptional defect caused by a mutation in another component of the transcription complex. We thus extended our analysis of the effect of the CURE sequence on activation of transcription from the CUP1 UAS–promoter region to a yeast strain deleted for Srb5 (srb5Δ). As in the case of GAL11, SRB5 is not an essential gene and therefore it is possible to use similar experimental conditions to test its requirement in gene transcription. Indeed, since a srb5Δ mutant strain is viable, no temperature shift is needed to inactivate the function of Srb5. We therefore constructed the mutant strain BLY29 with a complete deletion of the SRB5 locus (srb5Δ). We first measured Cu2+-induced β-galactosidase activities from the BLY29 strain transformed with the episomal reporter construct pSAL1::lacZ along with either the plasmid pBL12, expressing Srb5, or the empty parental plasmid YCplac33. The results of these experiments were similar to those obtained by monitoring the effect of deleting Gal11 on the expression of this reporter plasmid and indicated that Srb5 was also required for full activation of this episomal CUP1 UAS::lacZ reporter gene (data not shown). We subsequently integrated the pSAL1::lacZint and pSAL1–1441::lacZint reporter constructs at the LEU2 locus of BLY29 and, after Cu2+ induction, monitored expression of these reporter genes in the presence or absence of Srb5. The northern blot in Figure 5 shows that the negative effect of srb5Δ on CUP1 UAS-mediated activation of transcription persisted even when the CUP1 UAS::lacZ reporter gene was integrated at the LEU2 locus. Similarly to our previous observations with the gal11Δ mutant, the negative effect of srb5Δ on transcription was almost completely rescued by addition of the CURE sequence upstream of the CUP1 UAS without significantly affecting the maximal level of reporter gene expression observed in the presence of Srb5 protein (Fig. 5). We conclude that the ‘protective’ effect of the CURE sequence on the efficiency of transcriptional activation from the CUP1 UAS–promoter elements is not specific for the gal11Δ mutation, but can be extended to the deletion of at least one other component of the Srb/mediator complex, i.e. Srb5. Presence of the CURE sequence influences the local chromatin configuration of the CUP1 promoter region The results presented so far show that the CURE sequence, which is present upstream of the CUP1 UAS, is able to rescue impaired transcription in strains carrying mutations in the transcription complex components Gal11 and Srb5. To gain insights into how the CURE sequence might influence the activity of the downstream CUP1 UAS–promoter elements, we performed a series of chromatin assays in which we measured the degree of DNA accessibility to nucleases of the CUP1 UAS–promoter region in the presence or absence of the upstream CURE sequence. The BLY18 strain, which lacks the CURE sequence, and the BLY22 strain, which bears the CURE upstream of the integrated CUP1 UAS::lacZ construct, were analyzed for their chromatin organization at the sites of the LEU2-integrated CUP1 UAS reporter genes. This analysis was performed using two different nuclease hypersensitivity assays on the intact chromatin. We first analyzed the overall chromatin structure of the upstream regions of the CUP1 reporter genes by MNase digestion followed by gel electrophoresis and blotting. Indirect end-labeling of the blotted DNA fragments was utilized to identify chromatin-specific, and possibly CURE-dependent, MNase-hypersensitive sites. This assay was performed on chromatin obtained after the exposure of yeast cultures to copper ions to induce activation of the CUP1 enhancer–promoter region. Four relatively strong MNase-hypersensitive sites were found upstream of the BLY18 CUP1 UAS within a region of the reporter construct that lacked the CURE sequence. These sites were not, or only weakly, detectable within the CURE sequence at a similar distance upstream of the BLY22 CUP1 UAS (Fig. 6A, denoted by the open circles on the left). Three moderately hypersensitive sites appeared instead only within the CURE sequence (Fig. 6A, closed circles). The overall chromatin structure of the CUP1 UAS–promoter region did not appear to be affected by the presence or absence of the upstream CURE sequence with the exception of a possible change revealed by the diminished hypersensitivity to MNase of one site in the 5′-region of the CUP1 UAS (denoted by an asterisk in Fig. 6A) in the presence of the CURE. To more precisely map and quantitate changes in DNA accessibility of the CUP1 UAS–promoter region, chromatin from untreated (Fig. 6B) and Cu2+-treated BLY18 and BLY22 cells (Fig. 6C) was analyzed by restriction nuclease digestion. These experiments show that presence of the CURE sequence upstream of the CUP1 UAS region causes alterations in the accessibility of a set of restriction nuclease sites within the CUP1 UAS–promoter region. The pattern of this altered accessibility does not significantly change for chromatin prepared from Cu2+-treated (induced) as compared to untreated cells. However, as expected (26), the overall degree of accessibility clearly increased upon Cu2+ induction, with the exception of the MspA1I site that overlaps the Ace1-binding site (compare Fig. 6B with C). Access to this DNA site for MspA1I is most likely blocked by the presence of Ace1, which has been shown to bind its recognition sequences only upon Cu2+ induction. The most dramatic changes in DNA accessibility occurred at positions that lie within the 5′-portion of the cloned CUP1 UAS region, which became less accessible in the presence of the upstream CURE sequence (Fig. 6B and C, HaeII, NdeI and DraI). These results are in agreement with those from the MNase assays (asterisk in Fig. 6A). The differences in accessibility between the two promoter configurations (plus or minus the CURE) were smaller or absent for positions immediately upstream and downstream of the Ace1-binding sites. However, for those sites where differences were detectable, the accessibility trend changed from diminished accessibility to slightly higher accessibility in the presence of the CURE sequence (Fig. 6B and C, from SpeI to MunI). We conclude that the presence of the CURE sequence upstream of the CUP1 UAS did not generate the formation of an obvious, specific nucleosome pattern. However, this sequence had an influence on the chromatin structure rendering the DNA region surrounding the binding sites for the activators AceI and Hsf1 slightly more accessible, while the more distal 5′-sequences became less accessible to restriction nuclease activity. DISCUSSION Our principal results can be summarized as follows. Efficient activation of transcription of a reporter gene bearing the CUP1 UAS–promoter region isolated from its natural chromosomal context required the function of the transcription factors Gal11 and Srb5, two components of the Srb/mediator complex of the yeast RNA pol II transcriptional machinery. Insertion of the CURE upstream of the isolated CUP1 UAS–promoter region drastically reduced the dependence on these transcription factors for efficient activation of transcription without affecting the maximal levels of gene expression. Reciprocally, integration of this reporter gene at the CUP1 locus such as to reconstitute the CURE sequence upstream of the CUP1 UAS–promoter region also rendered activation of transcription much less sensitive to the absence of Gal11 and Srb5. Thus, the degree of dependence on these transcription factors (i.e. their utilization) for full activation from the CUP1 UAS is determined by the CURE sequence, which is naturally present upstream of each multimerized CUP1 gene. It has recently become clear that numerous components (transcription factors) of the pol II transcription complex are differentially utilized at different genes or classes of genes (5). The functional role of such differential utilization of transcription complex components has remained elusive. Young and collaborators have pointed out that the distinct expression signatures observed for a number of these components reveal a level of genome regulation that can be superimposed on that due to sequence-specific transcriptional activators and repressors (3,5). They have suggested that the expression of specific sets of genes might also be controlled by affecting the availability or function of specific components of the initiation complex, which, for example, may be targets of certain signal transduction pathways (5). It is also conceivable that some genes need to be rapidly and efficiently activated even in the absence of several transcription complex components, for example in response to drastic environmental changes that may impair the function of some transcription factors. CUP1 might be an example of such a gene. What determines the requirement and the role of any particular component of the transcription complex at any given gene? It has been suggested that different sequence-specific regulatory proteins (activators or repressors) may work by interacting with different transcription factors and thus determine, at least in part, which components of the transcription complex are required for the set of genes they specifically regulate (3). For example, recent studies by Lis and collaborators, which were aimed at analysis of the mechanism whereby activation of CUP1 transcription is insensitive to the depletion of Srb4 from the transcription complex, have shown that the CUP1 UAS-binding activators Ace1 and Hsf1 need to establish functional contacts with only a subset of transcription factors, which includes Rgr1 but not Srb4, to work optimally. Thus, mutated transcription complexes that lacked the components not required by Ace1, such as Srb4, could still be efficiently engaged in the CUP1 activation process, while a mutated complex that lacked Rgr1 only poorly responded to these specific activators (8). Core promoter sequences may also contribute to the differential requirements for components of the transcription complex. Indeed, Green and collaborators have shown that core promoter sequences, not the activator-binding elements, determine whether a defined gene requires the TFIID subunit TAF145 for activation of transcription in yeast (27). Similarly, genomic sequences that are not activator-binding sites appear to determine the dependence on the Swi/Snf complex for activation of the HO gene in yeast. Indeed, it has been shown that substitution of the binding sites for the HO-specific Swi5 activator by those for the Gal4 activator maintains the dependence on Swi/Snf, even though Gal4 does not usually require Swi/Snf for its function, as, for example, activation of the natural Gal4 target gene GAL1 is completely Swi/Snf independent (28). The results of our analysis of CUP1 activation show that the requirement for the transcription factors Gal11 and Srb5 at the CUP1 promoter is strongly dependent on the presence of the CURE sequence upstream of the CUP1 UAS. Thus, in this case the control for the requirement of these transcription factors at the CUP1 promoter is not exercised by the CUP1 UAS, but rather by a functionally novel sequence that lies outside the CUP1 UAS–promoter region. What is the mechanism by which the CURE sequence can drastically reduce the dependence on Gal11 and Srb5, and perhaps other transcription factors, for efficient activation from the CUP1 UAS–promoter region? Possible mechanisms implicate the presence of binding sites for transcription factors within the CURE sequence that may facilitate activation from the downstream CUP1 UAS by providing additional contacts to help recruit a crippled pol II machinery that lacks Gal11 or Srb5. In our experiments, such a putative helping effect of CURE-binding factors is not detectable in the presence of a wild-type pol II machinery, as the level of transcription in this case is not influenced by the CURE. It is also possible that the CURE sequence itself or specialized DNA-binding factors induce a transcriptionally more active chromatin structure in its vicinity. In agreement with this hypothesis, Grunstein and collaborators have shown that CUP1 and several stress-inducible genes, like HSP26, HSP70 (SSA3) and HSP70 (SSA4), can be activated upon nucleosome depletion in a UAS-independent fashion (29). Our analysis of the in vivo chromatin structure of the CUP1 UAS–promoter region showed that although there was no remarkable alteration in the overall nucleosomal pattern, the presence of the CURE caused changes in the DNA accessibility of this region (see Fig. 6). We are considering the possibility that the DNA accessibility pattern we observe near the CUP1 UAS might be a consequence of the recognition of sequences in the CURE region by trans-acting factors. The HMG1/2-like DNA architectural factors NHP6A/B are among the plausible candidates for such factors. Indeed, deletion of the genes encoding NHP6A/B was shown to negatively affect expression of a subset of genes, including CUP1 (30). These two proteins bind DNA without apparent sequence specificity and induce an overall bending of the helix through their HMG box DNA-binding domain (30,31). Thus, it is conceivable that the CURE sequence might provide a docking point for such architectural factors that would ultimately facilitate the accessibility of the CUP1 promoter to the transcription complex. In this work we consider the function of the CURE sequence with respect to its influence on CUP1 transcription. This region has so far been referred to as ORF YHR054C. The sequence of YHR054C shows strong similarity with the 3′-part of the neighboring YHR056C, from which it probably originated through duplication of this gene fragment. Despite the presence of an ORF, the ability of YHR054C to encode a functional protein product remains questionable (32). Some hypotheses regarding the necessity for co-amplification of YHR054C and CUP1 have been put forward. The amplification of YHR054C might lead to some selective advantage other than copper resistance, as copper-resistant strains seem to maintain multiple copies of the locus even in the absence of this particular selective pressure (33). Our results suggest that the sequence of YHR054C, namely the CURE, might provide a selective advantage through its role on CUP1 transcription rather than by giving rise to an independent gene product. Indeed, the CURE sequence positively acts on activation of CUP1 by turning a promoter that is dependent on transcription factors such as Gal11 and Srb5 into a promoter that is less sensitive to their deletion. We propose that the presence of the CURE within the CUP1 amplification unit sequence has been maintained through evolution to protect the stress-inducible CUP1 UAS–promoter elements against reduced activation efficiency that would otherwise result from crippled transcription complexes under stress conditions. ACKNOWLEDGEMENTS We thank Drs A. Auf der Maur, L. R. Martin, M. Petrascheck and J. F. Roth for their helpful comments on the manuscript and F. Ochsenbein for artwork. We are also grateful to Dr  A. Covarrubias for providing materials and to Dr M. Bodmer-Glavas for help with the chromatin analysis. This work was supported by research grants from the Helmut Horten Foundation and the Swiss National Research Fund. * To whom correspondence should be addressed at present address: ESBATech AG, Wagistrasse 21, CH-8952 Zürich-Schlieren, Switzerland. Tel: +41 1 635 3135; Fax: +41 1 635 6811; Email: [email protected] Present address:Laura Badi, F.Hoffmann-La Roche Ltd, Pharmaceuticals Division, CH-4002 Basel, Switzerland View largeDownload slide Figure 1. Lack of Gal11 significantly affects Cu2+-induced expression of an episomal CUP1 UAS::lacZ reporter gene but not that of the endogenous CUP1 genes. (A) Northern blot analysis of the relative levels of CUP1 and SNR6 transcripts isolated from JPY8 (gal11Δ) cells either expressing (Gal11+) or not (Gal11–) gal11 protein. SNR6 is transcribed by RNA pol III and is not affected by the lack of Gal11. (B) β-Galactosidase assay to measure the Cu2+-induced lacZ expression in JPY8 (gal11Δ) either containing Gal11 (+) or not (–). The experiment was performed in triplicate and the error bars indicate the standard deviation obtained for each set of samples. (C) Northern blot analysis of the same transformants; the arrow indicates the radioactive signal corresponding to the lacZ transcripts. The intensities of the 3.8 and 2.0 kb rRNA are shown as a measure of the constant amount of total RNA used in the northern blot assay. View largeDownload slide Figure 1. Lack of Gal11 significantly affects Cu2+-induced expression of an episomal CUP1 UAS::lacZ reporter gene but not that of the endogenous CUP1 genes. (A) Northern blot analysis of the relative levels of CUP1 and SNR6 transcripts isolated from JPY8 (gal11Δ) cells either expressing (Gal11+) or not (Gal11–) gal11 protein. SNR6 is transcribed by RNA pol III and is not affected by the lack of Gal11. (B) β-Galactosidase assay to measure the Cu2+-induced lacZ expression in JPY8 (gal11Δ) either containing Gal11 (+) or not (–). The experiment was performed in triplicate and the error bars indicate the standard deviation obtained for each set of samples. (C) Northern blot analysis of the same transformants; the arrow indicates the radioactive signal corresponding to the lacZ transcripts. The intensities of the 3.8 and 2.0 kb rRNA are shown as a measure of the constant amount of total RNA used in the northern blot assay. View largeDownload slide Figure 2. The effect of gal11Δ on activation of the CUP1 promoter is dependent on the chromosomal context of the integrated CUP1 UAS::lacZ reporter. (A) β-Galactosidase assay to measure lacZ expression in strains BLY91 and BLY97, either containing Gal11 (Gal11+) or not (Gal11–). The experiment was performed in triplicate and error bars indicate the standard deviation obtained for each set of samples. (B) S1 mapping of the lacZ transcripts under both non-inducing and inducing conditions (400 µM Cu2+). The free probe used in the protection assay is indicated by the letter P. Arrows indicate the position of the protected lacZ mRNA initiated at the two CUP1 start sites (+1 and +10). Note that the absolute levels of lacZ transcript are higher in BLY91 than in BLY97, reflecting the higher number of copies of the reporter gene integrated at the CUP1 locus, as shown in Figure 3. View largeDownload slide Figure 2. The effect of gal11Δ on activation of the CUP1 promoter is dependent on the chromosomal context of the integrated CUP1 UAS::lacZ reporter. (A) β-Galactosidase assay to measure lacZ expression in strains BLY91 and BLY97, either containing Gal11 (Gal11+) or not (Gal11–). The experiment was performed in triplicate and error bars indicate the standard deviation obtained for each set of samples. (B) S1 mapping of the lacZ transcripts under both non-inducing and inducing conditions (400 µM Cu2+). The free probe used in the protection assay is indicated by the letter P. Arrows indicate the position of the protected lacZ mRNA initiated at the two CUP1 start sites (+1 and +10). Note that the absolute levels of lacZ transcript are higher in BLY91 than in BLY97, reflecting the higher number of copies of the reporter gene integrated at the CUP1 locus, as shown in Figure 3. View largeDownload slide Figure 3. Organization of the CUP1 locus in the reporter strains BLY91 and BLY97. (A) Southern blot analysis; empty arrows indicate the bands originating from KpnI/ScaI double digested CUP1 repeat units which contain the CUP1 UAS::lacZ reporter in strains BLY91 and BLY97. The radiolabeled probe used for detection of the digestion products corresponds to the CUP1 UAS region [see (B)]. The filled arrow marks the 2.0 kb KpnI–KpnI band, which roughly corresponds to the CUP1 amplification unit, as shown by the digestion of genomic DNA from the JPY8 strain prior to integration of the CUP1 UAS::lacZ reporter (lane N). The size of the fragments is in kb. (B) Schematic representation of the multimerized CUP1 genomic locus and of the integrated CUP1 UAS::lacZ reporter constructs in BLY91 and BLY97. n represents the number of tandemly repeated units per haploid genome, with an extimated value of 15 for most laboratory strains (12). In the detailed structure of the generic (k) CUP1amplification unit (CUP1 AU), dashed lines between positions –210 and –105 mark the positions of the binding sites for Ace1 within the CUP1 UAS. Positions between –1469 and –394 delimit the CURE, while the CUP1downstream repeated element (CDRE) lies between positions +249 and +528. Numbers in bold indicate the two start sites. K, KpnI; S, ScaI; X, XbaI. Fragments which were detected in the Southern analysis in (A) are underlined and their length (in kb) is indicated below. In both BLY91 and BLY97 black arrowheads indicate the copies of the CUP1 UAS which are flanked by their natural CURE. In BLY91 an empty arrowhead marks the CUP1 UAS that is not linked to a CURE. View largeDownload slide Figure 3. Organization of the CUP1 locus in the reporter strains BLY91 and BLY97. (A) Southern blot analysis; empty arrows indicate the bands originating from KpnI/ScaI double digested CUP1 repeat units which contain the CUP1 UAS::lacZ reporter in strains BLY91 and BLY97. The radiolabeled probe used for detection of the digestion products corresponds to the CUP1 UAS region [see (B)]. The filled arrow marks the 2.0 kb KpnI–KpnI band, which roughly corresponds to the CUP1 amplification unit, as shown by the digestion of genomic DNA from the JPY8 strain prior to integration of the CUP1 UAS::lacZ reporter (lane N). The size of the fragments is in kb. (B) Schematic representation of the multimerized CUP1 genomic locus and of the integrated CUP1 UAS::lacZ reporter constructs in BLY91 and BLY97. n represents the number of tandemly repeated units per haploid genome, with an extimated value of 15 for most laboratory strains (12). In the detailed structure of the generic (k) CUP1amplification unit (CUP1 AU), dashed lines between positions –210 and –105 mark the positions of the binding sites for Ace1 within the CUP1 UAS. Positions between –1469 and –394 delimit the CURE, while the CUP1downstream repeated element (CDRE) lies between positions +249 and +528. Numbers in bold indicate the two start sites. K, KpnI; S, ScaI; X, XbaI. Fragments which were detected in the Southern analysis in (A) are underlined and their length (in kb) is indicated below. In both BLY91 and BLY97 black arrowheads indicate the copies of the CUP1 UAS which are flanked by their natural CURE. In BLY91 an empty arrowhead marks the CUP1 UAS that is not linked to a CURE. View largeDownload slide Figure 4. Insertion of the CURE 5′ of the CUP1 UAS::lacZ reporter confers insensitivity to gal11Δ. (A) β-Galactosidase assay to assess the level of Cu2+-induced lacZ expression in BLY18 and BLY22 either containing Gal11 (Gal11+) or not (Gal11–). The experiment was performed in triplicate and error bars indicate the standard deviation obtained for each set of samples. (B) Northern blot analysis; the arrow indicates the radioactive signal corresponding to the lacZ transcripts. The intensities of the 3.8 and 2.0 kb rRNAs are shown as a measure of the amount of total RNA considered in the northern blot analysis. View largeDownload slide Figure 4. Insertion of the CURE 5′ of the CUP1 UAS::lacZ reporter confers insensitivity to gal11Δ. (A) β-Galactosidase assay to assess the level of Cu2+-induced lacZ expression in BLY18 and BLY22 either containing Gal11 (Gal11+) or not (Gal11–). The experiment was performed in triplicate and error bars indicate the standard deviation obtained for each set of samples. (B) Northern blot analysis; the arrow indicates the radioactive signal corresponding to the lacZ transcripts. The intensities of the 3.8 and 2.0 kb rRNAs are shown as a measure of the amount of total RNA considered in the northern blot analysis. View largeDownload slide Figure 5. Low levels of CUP1 UAS activation in srb5Δ are rescued by addition of the CURE 5′ of the CUP1 UAS::lacZ reporter. Northern blot analysis showing the radioactive signals corresponding to lacZ and SNR6 transcripts detected for the reporter strains BLY30 (CUP1 UAS::lacZ) and BLY32 (CURE::lacZ), either expressing Srb5 (+) on not (–). View largeDownload slide Figure 5. Low levels of CUP1 UAS activation in srb5Δ are rescued by addition of the CURE 5′ of the CUP1 UAS::lacZ reporter. Northern blot analysis showing the radioactive signals corresponding to lacZ and SNR6 transcripts detected for the reporter strains BLY30 (CUP1 UAS::lacZ) and BLY32 (CURE::lacZ), either expressing Srb5 (+) on not (–). View large Download slide View large Download slide View large Download slide Figure 6. Alterations in local DNA accessibility within the CUP1 regulatory regions. (A) MNase analysis. A schematic representation of the ScaI (S)–PvuII (P) genomic regions considered for analysis is shown beside the autoradiogram. Lanes in which the ScaI–PvuII fragments contain or not the complete CURE sequence upstream of the CUP1 UAS are indicated by + and –, respectively. Open circle, bands detected primarily in the absence of CURE; closed circle, bands detected primarily in the presence of CURE; asterisk, band which is weaker relative to the lower band in the presence of CURE. The region in black corresponds to the lacZ reporter gene. The bent arrows mark the two transcription start sites. (B) Restriction endonuclease analysis before Cu2+ induction. The tested restriction enzymes that cut within the CUP1 UAS are indicated in the scheme at the top. The thick black line below this scheme indicates the probe used for Southern blotting. (C) Restriction endonuclease analysis after Cu2+ induction. In the graph below the autoradiograms, bars indicate accessibility of the CUP1 UAS in the absence (white bars) and presence (black bars) of the CURE upstream of the CUP1 UAS. Accessibility is expressed as a percentage of the signal from the band derived from digestion inside the CUP1 UAS (region indicated by the thick black arrow), relative to the total amount of ScaI–PvuII fragment considered for the analysis (R+) and (R). View large Download slide View large Download slide View large Download slide Figure 6. Alterations in local DNA accessibility within the CUP1 regulatory regions. (A) MNase analysis. A schematic representation of the ScaI (S)–PvuII (P) genomic regions considered for analysis is shown beside the autoradiogram. Lanes in which the ScaI–PvuII fragments contain or not the complete CURE sequence upstream of the CUP1 UAS are indicated by + and –, respectively. Open circle, bands detected primarily in the absence of CURE; closed circle, bands detected primarily in the presence of CURE; asterisk, band which is weaker relative to the lower band in the presence of CURE. The region in black corresponds to the lacZ reporter gene. The bent arrows mark the two transcription start sites. (B) Restriction endonuclease analysis before Cu2+ induction. The tested restriction enzymes that cut within the CUP1 UAS are indicated in the scheme at the top. The thick black line below this scheme indicates the probe used for Southern blotting. (C) Restriction endonuclease analysis after Cu2+ induction. In the graph below the autoradiograms, bars indicate accessibility of the CUP1 UAS in the absence (white bars) and presence (black bars) of the CURE upstream of the CUP1 UAS. Accessibility is expressed as a percentage of the signal from the band derived from digestion inside the CUP1 UAS (region indicated by the thick black arrow), relative to the total amount of ScaI–PvuII fragment considered for the analysis (R+) and (R). Table 1. Yeast strains used in this study Strain  Genotype  Integrated reporter plasmid  Integration (locus)  JPY5  MATα, lys2-202, ura3-52, his3-200, trp1-63, leu2-1  None    JPY8  MATα, lys2-202, ura3-52, his3-200, trp1-63, gal11Δ::TRP1, leu2-1  None    BLY91  MATα, lys2-202, ura3-52, his3-200, trp1-63, gal11Δ::TRP1  pSAL1::lacZint  CUP1 tandem  BLY97  MATα, lys2-202, ura3-52, his3-200, trp1-63, gal11Δ::TRP1  pSAL1::lacZint  CUP1 single 3′  BLY18  MATα, lys2-202, ura3-52, his3-200, trp1-63, gal11Δ::TRP1  pSAL1::lacZint  LEU2  BLY22  MATα, lys2-202, ura3-52, his3-200, trp1-63, gal11Δ::TRP1  pSAL1–1441::lacZint  LEU2  BLY29  MATα, lys2-202, ura3-52, his3-200, trp1-63, srb5Δ::TRP1, leu2-1  None    BLY30  MATα, lys2-202, ura3-52, his3-200, trp1-63, srb5Δ::TRP1  pSAL1::lacZint  LEU2  BLY32  MATα, lys2-202, ura3-52, his3-200, trp1-63, srb5Δ::TRP1  pSAL1–1441::lacZint  LEU2  Strain  Genotype  Integrated reporter plasmid  Integration (locus)  JPY5  MATα, lys2-202, ura3-52, his3-200, trp1-63, leu2-1  None    JPY8  MATα, lys2-202, ura3-52, his3-200, trp1-63, gal11Δ::TRP1, leu2-1  None    BLY91  MATα, lys2-202, ura3-52, his3-200, trp1-63, gal11Δ::TRP1  pSAL1::lacZint  CUP1 tandem  BLY97  MATα, lys2-202, ura3-52, his3-200, trp1-63, gal11Δ::TRP1  pSAL1::lacZint  CUP1 single 3′  BLY18  MATα, lys2-202, ura3-52, his3-200, trp1-63, gal11Δ::TRP1  pSAL1::lacZint  LEU2  BLY22  MATα, lys2-202, ura3-52, his3-200, trp1-63, gal11Δ::TRP1  pSAL1–1441::lacZint  LEU2  BLY29  MATα, lys2-202, ura3-52, his3-200, trp1-63, srb5Δ::TRP1, leu2-1  None    BLY30  MATα, lys2-202, ura3-52, his3-200, trp1-63, srb5Δ::TRP1  pSAL1::lacZint  LEU2  BLY32  MATα, lys2-202, ura3-52, his3-200, trp1-63, srb5Δ::TRP1  pSAL1–1441::lacZint  LEU2  View Large Table 2. Yeast plasmids   Name  Expressed gene/promoter element  Type  Marker  Expression vectors            pBL11  GAL11  ARS/CEN  URA3    pBL12  SRB5  ARS/CEN  URA3  lacZ reporter vectors            pSAL1::lacZ  CUP1 UAS–394  Episomal (ARS/CEN)  LEU2    pSAL1::lacZint  CUP1 UAS–394  Integrative  LEU2    pSAL1–1441::lacZint  CUP1 UAS–1441  Integrative  LEU2    Name  Expressed gene/promoter element  Type  Marker  Expression vectors            pBL11  GAL11  ARS/CEN  URA3    pBL12  SRB5  ARS/CEN  URA3  lacZ reporter vectors            pSAL1::lacZ  CUP1 UAS–394  Episomal (ARS/CEN)  LEU2    pSAL1::lacZint  CUP1 UAS–394  Integrative  LEU2    pSAL1–1441::lacZint  CUP1 UAS–1441  Integrative  LEU2  View Large References 1. Barberis,A. and Gaudreau,L. ( 1998) Recruitment of the RNA polymerase II holoenzyme and its implications in gene regulation. Biol. Chem. , 379, 1397–1405. Google Scholar 2. Ptashne,M. and Gann,A. ( 1997) Transcriptional activation by recruitment. Nature , 386, 569–577. 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van den Bosch, Michael;Zonneveld, José B. M.;Vreeken, Kees;de Vries, Femke A. T.;Lohman, Paul H. M.;Pastink, Albert

doi: 10.1093/nar/30.6.1316pmid: 11884628

Abstract In fission yeast two RAD52 homologs have been identified, rad22A+ and rad22B+. Two-hybrid experiments and GST pull-down assays revealed physical interaction between Rad22A and Rad22B, which is dependent on the N-terminal regions. Interaction with Rhp51 is dependent on the C-terminal parts of either protein. Both Rad22A and Rad22B also interact with RPA. The expression of rad22B+ in mitotically dividing cells is very low in comparison with rad22A+ but is strongly enhanced after induction of meiosis, in contrast to rad22A+. Rad22B mutant cells are not hypersensitive to DNA-damaging agents (X-rays, UV and cisplatin) and display normal levels of recombination. In these respects the Schizosaccharomyces pomberad22B mutant resembles the weak phenotype of vertebrate cells deficient for RAD52. Mutation of rad22A+ leads to severe sensitivity to DNA-damaging agents and to defects in recombination. In a rad22Arad22B double mutant a further increase in sensitivity to DNA-damaging agents and additional mitotic recombination defects were observed. The data presented here indicate that Rad22A and Rad22B have overlapping roles in repair and recombination, although specialized functions for each protein cannot be excluded. Received December 19, 2001; Revised and Accepted January 22, 2002. INTRODUCTION DNA double-strand breaks (DSBs) are among the most genotoxic DNA lesions and differ from most other types of damage in that both strands of the DNA double helix are affected. If not repaired or repaired incorrectly, DSBs may lead to gross chromosomal rearrangements, i.e. inversions, translocations and deletions, which are frequently found to be associated with carcinogenesis. Two main pathways to repair DSBs have been conserved during evolution: (i) homologous recombination (HR) and (ii) non-homologous end joining (NHEJ) (1). The relative importance of each mechanism in the repair of DSBs is dependent on the organism and phase of the cell cycle and may also be dependent on the nature of the DNA ends (1,2). Preservation of the genome by HR has been studied extensively in Saccharomyces cerevisiae and involves replication protein A (RPA), all three replicative DNA polymerases and the RAD52 group of genes (RAD50, RAD51, RAD52, RAD54, RAD55, RAD57, RAD59, MRE11, XRS2 and RDH54/TID1) (1,3). Although mutation of any of the genes in the RAD52 group results in sensitivity to DSB-inducing agents (i.e. ionizing radiation), rad52S.cerevisiae mutants display the most severe radiation sensitivity and defects in recombination (4). RAD52 is essential for meiotic recombination and almost all forms of mitotic recombination and gene conversion (4–6). Repair of a DSB flanked by repeated sequences can occur by a process called single-strand annealing (SSA), which can be considered as a subpathway of HR. SSA is dependent on RAD52 and partially on RAD59, especially when the DSB is flanked by short repeats (7,8). However, SSA does not require the presence of the RAD51, RAD54, RAD55 and RAD57 gene products (9). Telomere maintenance by a long tract gene conversion mechanism, known as break-induced replication, also relies on RAD52 and is independent of RAD51, RAD54, RAD55 and RAD57 (10–12). Consistent with the crucial role in SSA is the observation that Rad52 binds to single-stranded DNA (ssDNA) and that it mediates DNA strand annealing (13) in a reaction that is stimulated by RPA (14,15). After resection of the DSB, the second step in the conservative HR pathway is invasion of intact duplex DNA by one of the single-stranded ends and formation of D-loop DNA. This process of strand exchange requires the action of the RecA-related protein Rad51 (16). Recent investigations have shown that Rad52 and/or a heterodimer of Rad55 and Rad57 are involved in the nucleation of Rad51 onto ssDNA tails and promote strand exchange between the Rad51 nucleoprotein filament and the undamaged duplex DNA (17–20). The Rad52 protein, or the heterodimer of Rad55 and Rad57, stimulates the efficiency of these processes probably by overcoming the inhibitory effect that RPA has on DNA strand exchange. In agreement with these observations are findings that Rad52 interacts with Rad51 as well as with RPA (21–23). The biochemical activities of S.cerevisiae Rad52 are conserved in evolution. As in S.cerevisiae, Rad52 in man interacts with HsRad51 and HsRPA and stimulates the pairing and strand exchange activities of HsRad51 (24–27). Both Rad52 from S.cerevisiae and human Rad52 form multimeric ring structures that can bind to single- and double-stranded DNA ends (14,28). The importance of HR in the repair of DSBs in mammalian cells has been demonstrated by assays based on the repair of a site-specific DSB flanked by direct repeats. Repair by HR occurred in 30–50% of the molecules recovered (29). Moreover, inactivation of RAD54 in mouse ES cells and Drosophila melanogaster leads to reduced resistance to X-rays and defects in recombination (30,31). Mouse embryos homozygous for mutations in RAD51, RAD50 or MRE11 or in one of the RAD51 paralogs (XRCC2, RAD51L1 or RAD51L3) die during early embryonic development, implying an essential role in cell proliferation (32–38). In striking contrast to the severe phenotype of mutant rad52S.cerevisiae cells are the mild phenotypes of RAD52–/– mouse ES and chicken DT40 cells. Mouse and chicken cells deficient for RAD52 are not hypersensitive to DSB-inducing agents and show only subtle defects in HR (39,40). Moreover, deletion of MmRAD52 does not affect viability, fertility or lymphocyte development (40). An explanation for these observations could be the presence of additional genes functionally related to RAD52. Evidence for such a functional redundancy has been provided by the identification of the RAD59 gene in S.cerevisiae and Klyveromyces lactis (41,42). The Rad59 proteins of both S.cerevisiae and K.lactis display sequence homology to the N‐terminal domain of Rad52 protein, but lack the C-terminal part which is required for the interaction with Rad51. Deletion of RAD59 leads in both yeast species to a moderate X-ray sensitivity (41,42). Moreover, in S.cerevisiae it has been shown that a rad59 mutation can be partially complemented by overexpression of RAD52, suggesting that the functions of both proteins overlap (41). A set of two RAD52 homologs, rad22A+ and rad22B+, has also been isolated from the distantly related fission yeast Schizosaccharomyces pombe (43–45). Mutant rad22A cells are sensitive to DSB-inducing agents and show defects in the ability to perform homologous plasmid integration (45). Inactivation of rad22B+ does not lead to X-ray hypersensitivity, and homologous integration of linear DNA is not affected (45). Deletion of rad22B+ in a rad22A-deficient background, however, resulted in a further increase in radiation sensitivity. Overexpression of the rad22B+ gene in rad22A mutant cells partially suppresses the hypersensitivity to X‐rays, suggesting functional overlap between the two genes. Here we report that Rad22A and Rad22B can interact with each other. Both proteins also show physical interaction with Rhp51, the homolog of Rad51 in S.pombe, and RPA. In addition, we have investigated the S.pomberad22A, rad22B and rad22Arad22B mutant phenotypes in more detail, using mitotic and meiotic recombination assays. MATERIALS AND METHODS Strains and growth conditions The S.pombe strains used in this study are listed in Table 1. The media have been described elsewhere (46). Cells were grown in rich medium (YE) or YNB minimal medium [0.67% Yeast Nitrogen Base (Difco), 2% glucose]. For recombination assays Eagle’s minimal medium (EMM) was used. Appropriate amino acids were added to a final concentration of 100 mg/l. Geneticin selection was performed using YE medium containing 100 mg/l G418 (Gibco). Two-hybrid analysis The single copy two-hybrid vectors pPC97, carrying the Gal4 DNA-binding domain, and pPC86, carrying the Gal4 activation domain, have been described elsewhere (47). Gene fragments containing 5′ SalI and 3′ BamHI restriction sites were obtained by PCR using the Expand High Fidelity PCR System (Roche). The fragments obtained were first cloned in the Zero Blunt vector (Invitrogen) and verified by sequencing before cloning into the appropriate restriction sites of pPC97 and pPC86. Two-hybrid studies were performed using S.cerevisiae strain Y190 (48). Transformants were selected for tryptophan and leucine prototrophy on YNB medium containing 30 mg/l adenine. Protein–protein interactions were detected by a filter assay for β-galactosidase activity in permeabilized cells. At least 12 individual colonies from two independent transformations were tested. GST pull-down Protein–protein interactions were also investigated by GST pull-down experiments. DNA fragments were isolated as SalI–NotI fragments from two-hybrid clones and inserted into SalI + NotI-digested pGEX-6P-2 and pMV2-HA. The pMV2-HA vector contains the T7 promotor and can be used for the synthesis of HA-tagged fusion proteins in vitro. Glutathione S‐transferase (GST) fusion proteins were produced in Escherichia coli strain BL21 and bound to glutathione–Sepharose 4B beads essentially as described by the manufacturer (Pharmacia Biotech). Sepharose beads were stored at 4°C in phosphate-buffered saline, pH 7.4, containing 1% (v/v) Triton X-100 and the amount of protein immobilized was determined by SDS–PAGE. For the generation of 35S-labeled proteins, pMV2-HA clones were used as template in an in vitro coupled transcription/translation system (TNT System; Promega) in the presence of [35S]methionine. Binding assays were carried out for 60 min at 4°C in 500 µl of binding buffer (50 mM Tris–HCl pH 7.4, 250 mM NaCl, 5 mM EDTA, 50 mM NaF, 0.2% Nonidet P-40) containing 15 µl of 35S-labeled protein and 2.5 µg GST fusion protein loaded on Sepharose beads. The beads were washed five times with washing buffer (50 mM Tris–HCl pH 7.4, 500 mM NaCl, 5 mM EDTA, 50 mM NaF, 0.2% Nonidet P-40). If necessary, more stringent washes were performed in buffer containing 750 mM NaCl. Bound proteins were recovered by heating in 1× Laemmli buffer for 5 min, analyzed by PAGE and visualized by autoradiography. Protein analysis The diploid strain GP338 (see Table 1) was used for meiotic induction experiments essentially as described by Lin et al. (49). Cells were grown at 25°C in liquid EMM supplemented with adenine to OD600 = 0.3. The temperature was raised to 34°C to induce meiosis (49). At different time points, 10 ml samples were withdrawn and cells were harvested by centrifugation, washed once in sterile water and once with 20% trichloroacetic acid (TCA) and resuspended in 200 µl of 5% TCA. An equal volume of glass beads was added and the cells were lysed by vortexing. After addition of 400 µl of 5% TCA, the bottom of the tube was punctured and inserted into another test tube. The suspension was clarified by centrifugation for 5 min at 4000 r.p.m. To each sample was added 200 µl of 1× Western Loading Buffer (1× Western Loading Buffer = 1 vol 4× sample buffer, 2 vol 250 mM Tris–HCl pH 8.5, 1 vol H2O; 4× sample buffer = 250 mM Tris–HCl pH 6.8, 8% SDS, 20% glycerol, 20% 2-mercaptoethanol, 0.4% Bromophenol blue). Proteins were fractionated on 8% SDS–polyacrylamide gels and transferred to nitrocellulose. Anti-Rad22A polyclonal antibodies were used at 1:7000, anti-Rad22B antibodies at 1:5000 and anti-α-tubulin (Sigma) at 1:2000 dilution. Protein–antibody complexes were visualized by chemiluminescence using the Amersham ECL system. Inactivation of genes PCR-mediated deletion constructs were prepared in order to inactivate the rhp51+, rad22A+, rad22B+ and rhp54+ genes by replacing the ORF of these genes with the kanr gene that confers resistance to the aminoglycoside antibiotic G418. Synthesis of targeting fragments to delete the rad22A+ and rad22B+ genes was carried out using the following primer combinations and plasmid pUG6 (50) as template: rad22A+ sense, 5′-CTGCTTCATATAAGCTAGAAGGGATGTCTTTTGAGCAAAAACAGCATGTAGCATCAGAAGACCAGGGCCATTTT-AACCAGCTGAAGCTTCGTACGCTGC; rad22A+ antisense, 5′-AATCATTAGTCATAAAACAGAAAATACTTGGTAAAAA-ACAAGTTGCCAATCATCACATTTTGCCTCATTACTTTTATGCATAGGCCACTAGTGGATCTG; rad22B+ sense, 5′-GTGGCGAAAGACGCGTATAAAAAACCCATTATTCTTC-TTTTAACATTCCTTTAAATTAAGATCCCCAAAACATTTTCAAACAGCTGAAGCTTCGTACGC; rad22B+ antisense, 5′-GTTGTACAGCAATTTATTTCGCAAGCAGTTTAAAAGT-TTATACCATTGAACGTTTCACTCAATTTTATTTTTTTTATGCATAGGCCACTAGTGGATCTG. The nucleotide sequences underlined overlap with the template DNA. The following primers in combination with plasmid pFA6a-kanMX4 (51) were used to inactivate the rhp51+ and rhp54+ genes: rhp51+ sense, 5′-CGATTACAACTGGATCAAAGCAACTAGACACTTTGTTACAAGGAGGGGTTGAGACTGGAAGCATT-ACTGAATTATTTGGTGCGTACGCTGCAGGTCGAC; rhp51+ antisense, 5′-TTGGTTTTTTGGGATCAGGATTAAAGGAAATGCCATCCACTTGGGCGACCACCTGATTAGTAA-TGACAACAGCAATACCGATCGATGAATTCGAGCTCG; rhp54+ sense, 5′-TTTAGTGAAGAGTAACAATGGTGCGTAATTTAAAGAACGCGACACGCTATAATAGTAATAGAATTTAATTTATATATACATCGTACGCTGCAGGTCGAC; rhp54+ antisense, 5′-TTGCCTTGAACCTAACAAACCAGGATTCGCAAAATTTAACAACGAAAAGTATTCGCTTAGATCATTTTGAATAGGGGTACATCGATGAATTCGAGCTCG. The nucleotide sequences underlined overlap with the template DNA. PCR synthesis of short flanking homology disruption cassettes was carried out using the Expand High Fidelity PCR System (Roche) according to the instructions of the manufacturer. Several PCR reactions were pooled to yield ∼5 µg DNA, which was transformed into competent S.pombe cells according to the lithium acetate method as described by Keeney and Boeke (52). G418-resistant transformants were screened by diagnostic PCR to verify whether the disruption cassette had integrated at the correct position in the genome (data not shown). Details of the additional primers used for this purpose are available on request. Intrachromosomal recombination The intrachromosomal recombination substrate consists of two ade6 heteroalleles (ade6-M26 and ade6-L469) separated by plasmid sequences and a ura4+ gene (see Fig. 4) and has been described in detail by Osman et al. (53). Two classes of ade6+ recombinants can be obtained: Ade+ Ura+ conversion types and Ade+ Ura– deletion types (see Fig. 4). The assays for spontaneous direct repeat recombination were performed by fluctuation tests as described previously (53). For each experiment at least six independent colonies were used and each assay was repeated at least twice. Thus a total of at least 12 colonies per strain were analyzed. For the determination of spontaneous recombination frequencies, strains were grown on EMM containing 250 mg/l adenine, 100 mg/l uracil and 100 mg/l leucine. After 4 days growth, single colonies were analyzed by differential plating. Less diluted samples were plated on EMM lacking adenine (to select for Ade+ recombinants) but containing 100 mg/l uracil. Cell titers were determined by plating appropriate dilutions on complete medium. After 3–4 days growth, cell titer and number of Ade+ recombinants were determined. To determine the proportion of deletion versus conversion type events, Ade+ recombinants were replica-plated onto minimal medium lacking adenine and uracil, and medium lacking adenine but containing uracil. Homologous integration of linear DNA Recombination as measured by the integration of linear plasmid DNA at the leu1.32 locus was carried out as described previously (45,54). Aliquots of competent wild-type and isogenic mutant strains were transformed with 5 µg NdeI-linearized pJK148 according to the lithium acetate method as described by Keeney and Boeke (52). To determine the uptake of DNA, an aliquot of competent cells of each strain was transformed with 5 µg pREP1. leu1+ transformants were selected on EMM containing adenine and uracil. The recombination frequency of each strain was calculated by dividing the number of leu1+ transformants of pJK148-transformed cells by the number of leu1+ transformants of pREP1-transformed cells. Meiotic recombination Meiotic recombination frequencies between mutant alleles of ade6 (ade6-M26 and ade6-52) and between ade6 and arg1 and arg1 and ura4, respectively, were determined by random spore analysis. Crosses were performed on low nitrogen plates as described before (45). Dilution series of spores were plated on YE plates to determine the total number of spores. Less diluted samples were plated on YNB plates containing appropriate nutrients to select for recombinants. Recombinants were replica-plated onto sporulation medium and stained with iodine vapor after 4 days growth to detect diploid cells. The frequency of diploid cells (a diploid cell may contain two complementing chromosomes and can therefore be mistakenly counted as a recombinant) was extremely low in the experiments presented here. Therefore, the results given in Table 4 did not require correction. RESULTS Interactions between Rad22A, Rad22B, Rhp51 and Rhp54 In S.cerevisiae and mammals, physical interactions between Rad52 and Rad51 and between Rad52 and RPA have been detected (21–26). These and other studies suggested the presence of one or more protein complexes in which Rad52 resides with other members of the RAD52 group. To analyze possible interactions of Rad22A and Rad22B with other members of the RAD52 group in S.pombe, two-hybrid studies and GST pull-down experiments were performed. To reduce artificial interactions, single copy expression vectors were used for the two-hybrid experiments (47). When multicopy vectors are used, expression of rhp51+ in S.cerevisiae is toxic. All genes and gene fragments were tested as pairwise combinations of DNA-binding domain and activator fusions. Results were confirmed by reciprocal combinations of binding domain and activator fusions and are summarized in Figure 1. Autonomous lacZ activation was not observed for the constructs used in the experiments mentioned here. As may be expected on the basis of the results obtained with Rad52 homologs from S.cerevisiae and mammals, Rad22A from S.pombe is able to interact with Rhp51. A similar interaction was seen between Rad22B and Rhp51. The two-hybrid studies also indicated association between both Rad52 homologs from S.pombe, Rad22A and Rad22B. Rhp51, as well as Rad22A and Rad22B, showed self-interaction (Fig. 1). The self-interaction of Rad22A and Rad22B is compatible with the formation of ring structures, as has been reported for S.cerevisiae Rad52 and human Rad52 (14,28). Rad51 and its homolog in higher eukaryotes form a nucleoprotein filament on ssDNA (16,55,56). Similar properties are expected for Rhp51 and would explain the auto-interaction of this protein. The interaction seen between Rhp51 and Rhp54 is conserved in evolution, as similar observations have been made for Rad51 and Rad54 from S.cerevisiae and man (57–59). The two-hybrid experiments did not reveal an interaction between Rhp54 and Rad22A or Rad22B (Fig. 1). Interactions were also observed between Rad22B and Rhp57, between Rad22B and Ssb1 (the large subunit of S.pombe RPA) and between Rad22A and Ssb1, respectively (data not shown). Two-hybrid and GST pull-down experiments to identify possible interactions between Rad22A and Rhp57 were not conclusive. To localize the regions that mediate the interaction of Rad22A with Rad22B and Rhp51 and of Rad22B with Rad22A and Rhp51, respectively, N- and C-terminal deletion constructs were generated by PCR and cloned in pPC97 and pPC86. As shown in Figure 1, the truncated Rad22A-a protein, containing amino acids 1–167, is sufficient for binding to Rad22B and for self-interaction. Residues 259–469 of Rad22A form a complex with Rhp51. The N-terminal part of Rad22B (containing amino acids 1–158) is also sufficient for self-interaction and for interaction with Rad22A. For association with Rhp51, residues 233–371 of Rad22B are sufficient. The interactions as observed by two-hybrid analysis were confirmed by an in vitro binding assay (GST pull-down). Full-length Rad22A and Rad22B as well as C-terminal constructs were expressed in E.coli as GST fusion proteins. Proteins from bacterial lysates were bound to glutathione–Sepharose beads and incubated with [35S]methionine-labeled Rad22B-a, Rad22A-b or Rhp51. Glutathione–Sepharose beads containing GST protein were used as a control in all experiments. After washing, associated proteins were analyzed by gel electrophoresis. The results presented in Figure 2A and B indicate that the N-terminal domains of Rad22B and Rad22A are sufficient for interaction with Rad22A and Rad22B, respectively, and for self-interaction. Association with Rhp51 can be detected using full-length Rad22B and Rad22A and the C-terminal fragments of Rad22B and Rad22A (Fig. 2C and D). Rad22A and Rad22B protein levels In order to understand the phenotype of rad22B mutant cells and the mitotic and meiotic properties of rad22A cells, we investigated the transcript and protein levels of Rad22A and Rad22B. Whole cell extracts from mitotically growing cells and cells induced to undergo meiosis were prepared as described in Materials and Methods. As shown in Figure 3, expression of Rad22B protein was barely detectable in mitotically growing cells (0 h) and at 1–3 h after meiosis induction. At later stages in meiosis Rad22B protein can be detected. Rad22B protein is most abundant at 5 and 6 h after induction of meiosis. At later time points the level of Rad22B protein is reduced (Fig. 3). In rad22A mutant cells, expression of Rad22B is not increased relative to wild-type cells (data not shown). After probing with anti-Rad22B, the membrane was stripped and reprobed with anti-Rad22A. In contrast to Rad22B, Rad22A protein can be detected in both vegetatively growing cells and in cells induced to undergo meiosis. The level of Rad22A protein is not up-regulated after induction of meiosis (Fig. 3). Analysis of the transcript levels of rad22B+ by northern blot analysis also showed a strong increase in the RNA level of rad22B+ 5 h after induction of meiosis (data not shown). Transcripts of rad22A+ were easily detected in vegetative cells and a slight up-regulation in the RNA level was observed after induction of meiosis (data not shown). These observations indicate that Rad22A is more abundant than Rad22B in mitotically growing cells. Although not detectable by northern blot hybridization, the presence of rad22B+ RNA in mitotically growing cells was shown previously by RT–PCR (45). Recombination in vegetative cells Strains bearing direct repeats of ade6 heteroalleles, separated by plasmid sequences and a functional ura4+ gene (Fig. 4), were used to study spontaneous intrachromosomal recombination in mitotic cells as measured by the recovery of Ade+ recombinants. As indicated in Figure 4, two main classes of Ade+ recombinants can be distinguished: Ade+ Ura+ conversion types and Ade+ Ura– deletion types. Both types of events can occur within a single chromatid (intrachromosomal), but may also be due to recombination between sister chromatids after replication of the DNA. A gene conversion event not associated with crossing-over will result in an intact ade6+ gene and retention of the ura4+ gene. A similar outcome results by conservative one-sided invasion. On the other hand, possible mechanisms resulting in deletion type events are SSA, non-conservative one-sided invasion, gene conversion associated with crossing-over and unequal crossing-over between sister chromatids. In wild-type SL1 cells, spontaneous Ade+ recombinants arose at an average frequency of 3 per 104 viable cells. As indicated in Table 2, 52% of the recombinants were conversion type and the remainder deletion type. Next we determined the effects of several RAD52 group mutations on spontaneous direct repeat recombination. For rad22A, rad22B and rad22Arad22B mutant cells, no significant difference (P > 0.05) in the frequency of Ade+ recombinants (Table 2) was observed. However, a highly significant shift to deletion type events was observed in rad22A and rad22Arad22B mutant cells. About 84% of the Ade+ recombinants arose by a deletion type event in rad22A mutant cells. A preference for deletion type events in direct repeat recombination was also observed in rad22Arad22B mutant cells, as 89% of the recombinants arose by such a mechanism (Table 2). In rad22B mutant cells the distribution of recombination events was not different from that in wild-type cells (Table 2). The distribution of recombination events differs significantly between rad22A and rad22B mutant cells and also differs between rad22Arad22B and rad22B mutant cells (P < 0.01). No significant shift in conversion versus deletion types of Ade+ recovery is observed between rad22A and rad22Arad22B mutant cells. As indicated in Table 2, both rhp51 and rhp54 mutant strains exhibit a strong hyper-recombination phenotype. In both mutants recombination occurred primarily via a deletion type event. These latter observations are in agreement with previous S.pombe and S.cerevisiae studies using direct repeat recombination assays (60,61). Homologous recombination in the rad22Arad22B mutant was also studied by measuring the efficiency of integration of linear pJK148DNA at the leu1.32 locus (Table 3). The number of colonies obtained after transformation with linear pJK148 plasmid DNA was related to the number of pREP1 transformants. As reported previously, no reduction was seen in the pJK148/pREP1 ratio in the rad22B mutant and a 13-fold reduction was observed in the rad22A mutant (45). Inactivation of both rad22A+ and rad22B+ affected genomic integration of linear DNA at least 100-fold (Table 3). These results indicate that rad22B+ has the ability to contribute to HR, especially in the absence of rad22A+. Meiotic recombination In a previous study we showed that intragenic meiotic recombination at the ade6 locus, between ade6-M216 and ade6-469, was slightly affected in rad22A and rad22B mutant strains (45). The analysis of meiotic recombination was extended by measuring recombination at the intragenic interval between ade6-M26 and ade6-52 and by determining recombination at the intergenic intervals arg1–ade6 and arg1–ura4. As shown in Table 4, meiotic recombination in the rad22A, rad22B and rad22Arad22B mutant strains was only slightly affected at the ade6 locus. Recombination at ade6 was also studied in rhp51 and rhp54 mutant strains. Meiotic recombination was decreased ∼2-fold in the rhp51 mutant, whereas no reduction in intragenic meiotic recombination at ade6 was observed in the rhp54 mutant strain (data not shown). Intergenic meiotic recombination was determined at different genetic intervals on chromosome III between mutant alleles of arg1 and ade6 and between mutant alleles of arg1 and ura4. At both intervals the rad22B mutant showed a reproducible 1.5-fold increase in recombination (Table 4). Intergenic recombination at both intervals was reduced in the rad22A mutant strain (Table 4). Surprisingly, inactivation of rad22B+ increased the intergenic recombination frequencies of a rad22A mutant strain to the level of the rad22B single mutant (Table 4). DISCUSSION The role of Rad22A and Rad22B in vegetative cells The RAD52 homologs rad22A+ and rad22B+ from the fission yeast S.pombe show strong conservation at both the nucleotide and amino acid levels. Despite the similarities in sequence, S.pombe cells inactivated for rad22A+ show severe hypersensitivity to DNA-damaging agents (X-rays, bleomycin and UV light) and display defects in the ability to perform HR as measured by the integration of plasmid DNA (45). On the other hand, cells inactivated for rad22B+ are not hypersensitive to these DNA-damaging agents and are proficient in the ability to perform HR (44,45). Similar observations were made using cisplatin, which induces interstrand DNA crosslinks. In general, repair of these lesions also depends on HR. The rad22AS.pombe mutant displays increased sensitivity to cisplatin, however, the rad22B mutant is not hypersensitive to this compound (data not shown). In the rad22Arad22B double mutant the hypersensitivity is further increased (data not shown). Apparently, Rad22B is not required for the repair of a wide variety of DNA lesions if Rad22A is present. One explanation might be the low level of Rad22B in mitotically dividing cells (Fig. 3). The enhanced hypersensitivity of rad22Arad22B double mutant cells indicates that Rad22B functionally overlaps with Rad22A. The partial alleviation of the severe rad22A mutant phenotype by overexpression of rad22B+ is in agreement with this assumption (45). In addition, Rad22B might be required for the repair of specific types of DNA lesions. Very similar observations were made when HR was studied by integration of plasmid DNA. In the rad22Arad22B double mutant the level of recombination is further reduced in comparison with the rad22A single mutant (at least 7-fold; see Table 3). As for the repair of exogenously inflicted DNA damage, Rad22B functionally overlaps with Rad22A in this type of recombination. HR was also studied using a substrate containing direct repeats of two ade6 heteroalleles (Fig. 4). In the rad22 single and double mutant cells, recovery of Ade+ cells was not significantly different from that in wild-type cells (Table 2). However, mostly deletion type events were recovered in rad22A and rad22Arad22B mutant cells. A similar system has been used by McDonald and Rothstein (61) to study HR in S.cerevisiae cells deficient for RAD52. A slight reduction in the recovery of Ade+ colonies was observed in the S.cerevisiaerad52 mutant (61). As observed for S.pomberad22A and rad22Arad22B mutant cells, in rad52 mutant cells recombinants arose mainly by deletion type events. Apparently, rad22A and rad22Arad22B mutant cells can survive spontaneous DSBs by Rad22A/Rad22B-independent subpathways of recombination, which lead to deletions. The results of the protein–protein interaction studies are in agreement with a functional overlap between Rad22A and Rad22B. Both proteins show interaction with Rhp51 and with the large subunit of RPA. Also, in S.cerevisiae and higher eukaryotes Rad52 protein is able to interact with Rad51 and with RPA (21–26). As biochemical studies have indicated, the ability of Rad52 to interact with RPA and with Rad51 is essential for Rad51-dependent pairing and strand exchange reactions, which are the first steps in HR (17,19,20). The Rad52 protein from S.cerevisiae and man is able to form multimeric ring structures (14,28). In S.pombe, similar structures may consist of either Rad22A or Rad22B, or both proteins. Biochemical studies using purified Rad22A and Rad22B may provide more detailed information on the similarities and possible differences between both proteins. Meiotic properties of Rad22A and Rad22B It has been shown that the formation of DSBs during early meiosis is essential to promote meiotic recombination in S.cerevisiae and S.pombe (62–64). In both organisms, DSB formation during meiosis requires the function of multiple gene products. For example, inactivation of the meiosis-specific endonuclease SPO11 in S.cerevisiae and its corresponding homolog in S.pombe (rec12+) affects DSB formation during meiosis and results in drastically reduced meiotic recombination frequencies (>100-fold reduction) (64,65). In the rad22A mutant, meiotic intra- and intergenic recombination is slightly reduced at the ade6 locus and the arg1–ade6 interval, respectively, but is more strongly reduced at the larger arg1–ura4 interval. Spore viability is 4-fold reduced in the rad22AS.pombe mutant (45). In the fraction of surviving spores DSBs are probably repaired by rad22A+-independent mechanisms which may depend on rad22B+. The meiotic phenotype of S.cerevisiaerad52 mutant cells is much more severe than that of rad22AS.pombe cells. Meiotic recombination is at least 100-fold reduced and the production of viable spores is >200-fold decreased in the S.cerevisiaerad52 mutant (5). In contrast to rad22A+, expression of the rad22B+ gene is strongly regulated during meiosis. However, inactivation of rad22B+ does not affect spore viability and results in a very small increase in meiotic recombination at both intergenic intervals tested (Table 4). Surprisingly, the meiotic recombination frequency in the rad22Arad22B double mutant is comparable to the level of recombination in the rad22B single mutant. However, the viability of spores is dramatically reduced in the double mutant (45). The results show that residual repair of meiosis-specific DSBs is possible in the absence of Rad22A and Rad22B in the surviving fraction of spores. The level of recombination is hardly influenced in the surviving cells. In agreement with the meiotic recombination data and reduced spore viability are observations that meiotic DSBs are formed and accumulate in the rad22A and rad22Arad22B mutant strains (J.Young, C.Rubio and G.R.Smith, Fred Hutchinson Cancer Center, Seattle, WA, USA, personal communication). A similar phenotype has been observed for rhp51 mutants. Spore viability is drastically reduced, but the level of meiotic recombination is only slightly affected (54). Moreover, meiotic DSBs also accumulate in the rhp51 mutant strain (66). Rad52 mutant phenotypes The RAD52 gene from S.cerevisiae is required for nearly all (sub)pathways of recombination, as reviewed by Paques and Haber (3). The results presented here suggest that in S.pombe two proteins, Rad22A and Rad22B, perform the essential role that Rad52 has in most types of recombination in S.cerevisiae. However, minor Rad22A/Rad22B-independent mitotic and meiotic recombination pathways exist in S.pombe, explaining the less dramatic recombination defects of the double mutant in comparison with the S.cerevisiaerad52 mutant. Inactivation of rad22B+ hardly affects recombination and cell survival after exposure to DNA-damaging agents. In these respects, the rad22BS.pombe mutant resembles RAD52-deficient mouse ES and chicken DT40 cells. A second RAD52 homolog, which could be functionally related to rad22A+ in S.pombe, has not yet been identified in vertebrate cells. It cannot be excluded, however, that a second homolog is only functionally conserved and not structurally. Another possibility to explain the weak phenotype of RAD52-deficient mouse and chicken cells is functional complementation by other members of the RAD52 epistasis group. Rad51-dependent pairing and strand exchange reactions are stimulated by the addition of Rad52, and also by the addition of Rad55/Rad57, which are Rad51 paralogs (18). Recently, Fujimori et al. (67) reported that XRCC3–/–RAD52–/– double mutant DT40 cells could not be isolated, whereas either single mutant is viable. Possibly, the major contribution to assisting Rad51 in the formation of a nucleoprotein filament has shifted from Rad52 in S.cerevisiae towards the Rad51 paralogs in vertebrate cells. Therefore, it cannot be excluded that vertebrates have only one RAD52 homolog and that mutation of this gene is complemented by one (or more) of the RAD51 paralogs in higher eukaryotes. ACKNOWLEDGEMENTS The authors thank F. Osman for kindly providing the S.pombe SL1 strain, G. Smith for sharing unpublished data and sending the GP338 strain, J. Hegemann for providing the pUG6 vector, P. Philippsen for sending plasmid pFA6a-KANMX4, J. Dorsman for discussions and N. de Wind for critical reading of the manuscript. This work was supported by the Netherlands Organization for Scientific Research (NWO), grant 901-01-97. * To whom correspondence should be addressed. Tel: +31 71 5276152; Fax: +31 71 5276173; Email: [email protected] View largeDownload slide Figure 1. Schematic presentation of the protein–protein interactions investigated by two-hybrid analysis. All gene fragments were tested as DNA-binding fusions, as well as activator fusions. With the exception of the Rhp51–Rhp54 interaction, all the interactions were confirmed in the reciprocal orientation. Interactions were identified using qualitative β-galactosidase filter assays. +, positive interaction; –, no specific interaction; ND, not determined. View largeDownload slide Figure 1. Schematic presentation of the protein–protein interactions investigated by two-hybrid analysis. All gene fragments were tested as DNA-binding fusions, as well as activator fusions. With the exception of the Rhp51–Rhp54 interaction, all the interactions were confirmed in the reciprocal orientation. Interactions were identified using qualitative β-galactosidase filter assays. +, positive interaction; –, no specific interaction; ND, not determined. View largeDownload slide Figure 2.In vitro protein–protein interactions studied by GST pull-down assays. (A) An aliquot of 3 µl of 35S-labeled Rad22B-a (nucleotides 1–158) (lane 1); 15 µl of 35S-labeled Rad22B-a was mixed with glutathione–Sepharose beads loaded with 2.5 µg GST (lane 2), 2.5 µg GST::Rad22A (lane 3) or 2.5 µg GST::Rad22B (lane 4). (B) An aliquot of 3 µl of in vitro labeled Rad22A-b (nucleotides 1–258) (lane 1); 15 µl of 35S-labeled Rad22A-b was mixed with glutathione–Sepharose beads loaded with 2.5 µg GST (lane 2), 2.5 µg GST::Rad22A (lane 3) or 2.5 µg GST::Rad22B (lane 4). (C) An aliquot of 3 µl of in vitro labeled Rhp51 (lane 1); 15 µl of 35S-labeled Rhp51 was mixed with glutathione–Sepharose beads loaded with 2.5 µg GST (lane 2), 2.5 µg GST::Rad22B (lane 3), 2.5 µg GST::Rad22B-c (lane 4) or 2.5 µg GST::Rad22B-d (lane 5). (D) An aliquot of 3 µl of in vitro labeled Rhp51 (lane 1); 15 µl of 35S-labeled Rhp51 was mixed with glutathione–Sepharose beads loaded with 2.5 µg GST (lane 2), 2.5 µg GST::Rad22A (lane 3), 2.5 µg GST::Rad22A-c (lane 4) or 2.5 µg GST::Rad22A-d (lane 5). View largeDownload slide Figure 2.In vitro protein–protein interactions studied by GST pull-down assays. (A) An aliquot of 3 µl of 35S-labeled Rad22B-a (nucleotides 1–158) (lane 1); 15 µl of 35S-labeled Rad22B-a was mixed with glutathione–Sepharose beads loaded with 2.5 µg GST (lane 2), 2.5 µg GST::Rad22A (lane 3) or 2.5 µg GST::Rad22B (lane 4). (B) An aliquot of 3 µl of in vitro labeled Rad22A-b (nucleotides 1–258) (lane 1); 15 µl of 35S-labeled Rad22A-b was mixed with glutathione–Sepharose beads loaded with 2.5 µg GST (lane 2), 2.5 µg GST::Rad22A (lane 3) or 2.5 µg GST::Rad22B (lane 4). (C) An aliquot of 3 µl of in vitro labeled Rhp51 (lane 1); 15 µl of 35S-labeled Rhp51 was mixed with glutathione–Sepharose beads loaded with 2.5 µg GST (lane 2), 2.5 µg GST::Rad22B (lane 3), 2.5 µg GST::Rad22B-c (lane 4) or 2.5 µg GST::Rad22B-d (lane 5). (D) An aliquot of 3 µl of in vitro labeled Rhp51 (lane 1); 15 µl of 35S-labeled Rhp51 was mixed with glutathione–Sepharose beads loaded with 2.5 µg GST (lane 2), 2.5 µg GST::Rad22A (lane 3), 2.5 µg GST::Rad22A-c (lane 4) or 2.5 µg GST::Rad22A-d (lane 5). View largeDownload slide Figure 3. Relative expression of the Rad22A and Rad22B proteins in mitotically dividing cells (t = 0) and in cells induced to undergo meiosis (t = 1–8). To serve as a control for equivalent loading of proteins, blots were re-probed with an antibody raised against β-tubulin. View largeDownload slide Figure 3. Relative expression of the Rad22A and Rad22B proteins in mitotically dividing cells (t = 0) and in cells induced to undergo meiosis (t = 1–8). To serve as a control for equivalent loading of proteins, blots were re-probed with an antibody raised against β-tubulin. View largeDownload slide Figure 4. Schematic representation of the intrachromosomal recombination substrate and the two classes of Ade+ recombinants, conversion types and deletion types that can be distinguished. View largeDownload slide Figure 4. Schematic representation of the intrachromosomal recombination substrate and the two classes of Ade+ recombinants, conversion types and deletion types that can be distinguished. Table 1. Schizosaccharomyces pombe strains used in this study Strain  Genotype  RGL2  smt-0 ura4-D18 leu1-32 ade6-M26  RGL14  smt-0 rad22A:: ura4+ura4-D18 leu1-32 ade6-M26  RGL15  smt-0 rad22B::LEU2 ura4-D18 leu1-32 ade6-M26  RGL16  smt-0 rad22A::ura4+rad22B::LEU2 ura4-D18 leu1-32 ade6-M26  RGL17  h+rad22A::ura4+rad22B::LEU2 ura4-D18 leu1-32 ade6-M26  RGL20  h+ura4-D18 leu1-32 ade6-52  RGL21  h+rad22A::ura4+ura4-D18 leu1-32 ade6-52  RGL22  h+rad22B::LEU2 ura4-D18 leu1-32 ade6-52  RGL23  h+rad22A::ura4+rad22B::LEU2 ura4-D18 leu1-32 ade6-52  RGL26  h–leu1-32 arg1-14  RGL27  h+rad22A::loxKANMX4lox leu1-32 arg1.14  RGL28  h+rad22B::LEU2 leu1-32 arg1-14  RGL29  h+rad22A::loxKANMX4lox rad22B::LEU2 leu1-32 arg1-14  RGL30  h+leu1-32 ura4-595  RGL31  smt-0 rad22A::loxKANMX4lox ura4-595  RGL32  smt-0 rad22B::LEU2 leu1-32 ura4-595  RGL33  smt-0 rad22A::loxKANMX4lox rad22B::LEU2 leu1-32 ura4-595  RGL34  h+ura4-D18 leu1-32  RGL35  h+rad22A::ura4+ura4-D18 leu1-32  RGL36  h+rad22B::loxKANMX4lox ura4-D18 leu1-32  RGL37  h+rad22A::ura4+rad22B::loxKANMX4lox ura4-D18 leu1-32  SL1  h+ura4-D18 leu1-32 ade6-M26 int::pUC8/ura4/MATa/ade6-L469  RGL38  rad22A::loxKANMX4lox SL1  RGL39  rad22B::loxKANMX4lox SL1  RGL40  rad22A::loxKANMX4lox rad22B::LEU2 SL1  RGL41  rhp51::KANMX4 SL1  RGL42  rhp54::KANMX4 SL1  GP338  h–/h–ura4-294/+ +/leu1-32 ade6-M26/ade6-M210 + /arg1-2 pat1-114/pat1-114 end1-458/end1-458  Strain  Genotype  RGL2  smt-0 ura4-D18 leu1-32 ade6-M26  RGL14  smt-0 rad22A:: ura4+ura4-D18 leu1-32 ade6-M26  RGL15  smt-0 rad22B::LEU2 ura4-D18 leu1-32 ade6-M26  RGL16  smt-0 rad22A::ura4+rad22B::LEU2 ura4-D18 leu1-32 ade6-M26  RGL17  h+rad22A::ura4+rad22B::LEU2 ura4-D18 leu1-32 ade6-M26  RGL20  h+ura4-D18 leu1-32 ade6-52  RGL21  h+rad22A::ura4+ura4-D18 leu1-32 ade6-52  RGL22  h+rad22B::LEU2 ura4-D18 leu1-32 ade6-52  RGL23  h+rad22A::ura4+rad22B::LEU2 ura4-D18 leu1-32 ade6-52  RGL26  h–leu1-32 arg1-14  RGL27  h+rad22A::loxKANMX4lox leu1-32 arg1.14  RGL28  h+rad22B::LEU2 leu1-32 arg1-14  RGL29  h+rad22A::loxKANMX4lox rad22B::LEU2 leu1-32 arg1-14  RGL30  h+leu1-32 ura4-595  RGL31  smt-0 rad22A::loxKANMX4lox ura4-595  RGL32  smt-0 rad22B::LEU2 leu1-32 ura4-595  RGL33  smt-0 rad22A::loxKANMX4lox rad22B::LEU2 leu1-32 ura4-595  RGL34  h+ura4-D18 leu1-32  RGL35  h+rad22A::ura4+ura4-D18 leu1-32  RGL36  h+rad22B::loxKANMX4lox ura4-D18 leu1-32  RGL37  h+rad22A::ura4+rad22B::loxKANMX4lox ura4-D18 leu1-32  SL1  h+ura4-D18 leu1-32 ade6-M26 int::pUC8/ura4/MATa/ade6-L469  RGL38  rad22A::loxKANMX4lox SL1  RGL39  rad22B::loxKANMX4lox SL1  RGL40  rad22A::loxKANMX4lox rad22B::LEU2 SL1  RGL41  rhp51::KANMX4 SL1  RGL42  rhp54::KANMX4 SL1  GP338  h–/h–ura4-294/+ +/leu1-32 ade6-M26/ade6-M210 + /arg1-2 pat1-114/pat1-114 end1-458/end1-458  View Large Table 2. Spontaneous intrachromosomal mitotic recombination frequencies Strain  Total Ade+  P  Deletion types (%)  Conversion types (%)  P  Wild type   2.96 ± 1.16  –  48  52  –  rad22A   2.84 ± 1.35  >0.05  84  16  <0.01  rad22B   5.30 ± 3.18  >0.05  49  51  >0.5  rad22Arad22B   1.55 ± 0.51  >0.05  89  11  <0.01  rhp51  10.93 ± 4.36  <0.05  99   1  <0.01  rhp54  20.50 ± 6.66  <0.01  97   3  <0.01  Strain  Total Ade+  P  Deletion types (%)  Conversion types (%)  P  Wild type   2.96 ± 1.16  –  48  52  –  rad22A   2.84 ± 1.35  >0.05  84  16  <0.01  rad22B   5.30 ± 3.18  >0.05  49  51  >0.5  rad22Arad22B   1.55 ± 0.51  >0.05  89  11  <0.01  rhp51  10.93 ± 4.36  <0.05  99   1  <0.01  rhp54  20.50 ± 6.66  <0.01  97   3  <0.01  Recombination frequencies (per 104 viable cells ± SD) and percentages of deletion type versus conversion type were obtained as described in Materials and Methods. P values are from two-sample t-tests in which individual recombination frequencies and percentage deletion types of all colonies assayed were analyzed. P < 0.05 indicates a significant difference in recombination frequency (and percentage deletion type) for a given mutant strain, compared with the wild-type strain, at the 95% confidence level. P < 0.01 indicates a very significant (99% confidence interval) difference. View Large Table 3. Recombination as measured by the integration of plasmid DNA Strain  Leu+ transformantsa      pJK148/pREP1 ratio  Fold reduction  Wild-type  0.450  1  rad22A  0.036  13  rad22B  0.440  1  rad22Arad22B  0.003  >100  Strain  Leu+ transformantsa      pJK148/pREP1 ratio  Fold reduction  Wild-type  0.450  1  rad22A  0.036  13  rad22B  0.440  1  rad22Arad22B  0.003  >100  aThe reduction in the frequency of Leu+ transformants was calculated as described in Materials and Methods. View Large Table 4. Meiotic recombination frequencies Strain  Interval      Spore viabilityd    ade6-M26–ade6-52a  arg1-14– ade6-M26b  arg1-14– ura4-595c    Wild-type  75.1 ± 12.9  12.4 ± 2.1  15.9  92  rad22A  64.5 ± 10.8   7.9 ± 0.2   2.7  23  rad22B  70.6 ± 10.3  21.9 ± 5.3  23.2  88  rad22Arad22B  60.3 ± 2.1  20.0 ± 5.3  23.1   3  Strain  Interval      Spore viabilityd    ade6-M26–ade6-52a  arg1-14– ade6-M26b  arg1-14– ura4-595c    Wild-type  75.1 ± 12.9  12.4 ± 2.1  15.9  92  rad22A  64.5 ± 10.8   7.9 ± 0.2   2.7  23  rad22B  70.6 ± 10.3  21.9 ± 5.3  23.2  88  rad22Arad22B  60.3 ± 2.1  20.0 ± 5.3  23.1   3  aAverage recombination frequency per 104 cells of at least four independent crosses (± SD). bAverage recombination frequency per 102 cells of at least four independent crosses (± SD). cAverage recombination frequency per 102 cells of two independent crosses. dResults obtained previously (45). View Large References 1. Pastink,A., Eeken,J.C. and Lohman,P.H. 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Korn, Christian;Scholz, Sebastian Richard;Gimadutdinow, Oleg;Pingoud, Alfred;Meiss, Gregor

doi: 10.1093/nar/30.6.1325pmid: 11884629

Abstract The caspase-activated DNase (CAD) is involved in DNA degradation during apoptosis. Chemical modification of murine CAD with the lysine-specific reagent 2,4,6-trinitrobenzenesulphonic acid and the tyrosine-specific reagent N-acetylimidazole leads to inactivation of the nuclease, indicating that lysine and tyrosine residues are important for DNA cleavage by this enzyme. The presence of DNA or the inhibitor ICAD-L protects the enzyme from modification. Amino acid substitution in murine CAD of lysines and tyrosines conserved in CADs from five different species leads to variants with little if any catalytic activity, but unaltered DNA binding (K155Q, K301Q, K310Q, Y247F), with the exception of Y170F, which retains wild-type activity. Similarly, as observed for the previously characterised H242N, H263N, H308N and H313N variants, the newly introduced His→Asp/Glu or Arg exchanges lead to variants with <1% of wild-type activity, with two exceptions: H313R shows wild-type activity, and H308D at pH 5.0 exhibits ∼5% of wild-type activity at this pH. Y170F and H313R produce a specific pattern of fragments, different from wild-type CAD, which degrades DNA non-specifically. The recombinant nuclease variants produced in Escherichia coli were tested for their ability to form nucleolytically active oligomers. They did not show any significant deviation from the wild-type enzyme. Based on these and published data possible roles of the amino acid residues under investigation are discussed. Received December 14, 2001; Revised and Accepted January 17, 2002. INTRODUCTION Apoptosis in its final stages is characterised by nuclear DNA fragmentation (1). The major nucleolytic activity for the cell autonomous apoptotic DNA fragmentation in mammalian cells is the caspase-activated DNase (CAD), also termed DFF40 (DNA fragmentation factor 40 kDa subunit) (2–4). In non-apoptotic cells, this nuclease is bound to an inhibitory protein named ICAD-L/DFF45 (inhibitor of CAD large form/DNA fragmentation factor 45 kDa subunit), forming a heterodimeric complex known as DFF (DNA fragmentation factor). ICAD-L/DFF45 also serves as a specific chaperone for CAD/DFF40 and thus is required for the proper folding of a catalytically competent nuclease (5). Upon activation of caspases in the course of programmed cell death, ICAD-L/DFF45 is cleaved at two specific sites, releasing the nuclease from the DFF complex (2,6). Free CAD/DFF40 oligomerises into its active form, which degrades chromosomal DNA into nucleosomal units (7). Proteins such as histone H1, HMG1, HMG2 and topoisomerase II α bind to CAD/DFF40 and thereby stimulate its activity (7–9). A deletion analysis of CAD/DFF40 suggested that the catalytic centre is located in the C-terminal part of the enzyme (10,11), whereas structural and mutational studies revealed that the regulatory N-terminal domain (‘CAD-’ or ‘CIDE-N-domain’) is essential for a functional interaction of CAD/DFF40 with its chaperone and inhibitor ICAD-L/DFF45 (12–15). The C-terminal catalytic domain of CAD/DFF40 contains six conserved histidine residues, some of which, on the basis of chemical modification and site-directed mutagenesis studies, have been shown to be essential for DNA binding and/or cleavage by CAD/DFF40 (16,17). Besides the participation of histidine residues in DNA cleavage and binding by CAD/DFF40, which we have further explored in the present work, very little is known about the composition of the active site of this enzyme. In search of other functionally relevant amino acid residues and in the absence of structural information about the catalytic domain of CAD/DFF40, we have chemically modified recombinant CAD with the lysine-specific reagent 2,4,6-trinitrobenzenesulphonic acid (TNBS) and the tyrosine-specific reagent N-acetylimidazole (NAI). Our experiments show that lysine and tyrosine residues are essential for CAD/DFF40 activity. Guided by an amino acid sequence alignment of five homologous apoptotic nucleases, we have then replaced three conserved lysine residues and two conserved tyrosine residues in the catalytic domain of CAD by glutamine and phenylalanine, respectively. All variants were proficient in DNA binding and, with one exception, almost inactive in DNA cleavage. On the basis of these and previous results, possible roles of the amino acid residues under investigation in the mechanism of DNA binding and cleavage by CAD/DFF40 and the inhibition of CAD/DFF40 by ICAD-L/DFF45 are discussed. MATERIALS AND METHODS Expression of GST-mCAD/hICAD-L complex in Escherichia coli and preparation of free GST-mCAD GST-mCAD/hICAD-L complex and free GST-mCAD were produced as described previously (17). In brief, the GST-mCAD/hICAD-L complex was expressed in E.coli BL21Gold(DE3) cells harbouring plasmids pACET-DFF45 coding for wild-type human ICAD-L and pGEX-2T coding for wild-type GST-tagged murine CAD or the mutant versions, respectively, and purified by affinity chromatography on glutathione–Sepharose 4B beads. To obtain free GST-mCAD, the GST-mCAD/hICAD-L complex bound to the beads was incubated with recombinant caspase-3. After washing with buffer A (20 mM HEPES–NaOH pH 7.4, 100 mM NaCl, 1 mM EDTA) supplemented with 5 mM DTT, 10% glycerol, 0.01% Triton X-100 and 5 mM MgCl2, the free nuclease was eluted from the glutathione–Sepharose 4B beads and dialysed against buffer B (50 mM Na-phosphate, pH 7.5, 100 mM NaCl, 10% glycerol and 1 mM DTT). Protein concentrations were determined by UV spectroscopy, using a molar extinction coefficient calculated according to Pace et al. (18). Expression of mCAD in mammalian cells For expression of the mCAD/GST-hICAD-L complex in mammalian cells, the coding regions for wild-type mCAD and the variant Y170F were inserted into the vector pCS2-MT (19,20), and the coding region for a GST-hICAD-L/DFF45 fusion protein was inserted into the vector pCI (Promega). 293T human embryonic kidney cells were cultured in DMEM containing 10% fetal calf serum. Ten micrograms of each expression construct (pCI-GST-hICAD-L/DFF45 and pCS2-MT-mCAD) were used to co-transfect cells cultured in maxi-dishes using TransFast™ (Promega) transfection reagent according to the supplier’s recommendations. Forty-eight hours after transfection, cells were harvested and washed twice with PBS (1.5 ml/maxi-dish). After washing, cells were pelleted, resuspended and lysed by sonication in buffer A supplemented with 5 mM DTT and 10% glycerol. After sonication, Triton X-100 was added to a final concentration of 0.01%. Subsequently, the crude extract was centrifuged and the complex contained in the supernatant was bound to 75 µl of a suspension of glutathione–Sepharose 4B beads. Bound protein was washed twice with the same buffer as described above and resuspended in 175 µl of buffer A supplemented with 5 mM DTT, 10% glycerol and 0.01% Chaps {3-([3-cholamidopropyl]dimethylammonio)-1-propanesulfonate}. Free mCAD was eluted by treating the complex bound to the beads with 5 µl of a caspase-3 preparation for 1 h at ambient temperature. For DNA cleavage, 19 µl of eluted mCAD were incubated with 30 ng/µl assay solution of the plasmid pBSK-VDEX (New England Biolabs) for 15 min at 37°C in a total volume of 25 µl in buffer C (20 mM Tris–HCl, pH 7.4, 100 mM NaCl, 1 mM EDTA, 10 mM DTT, 5% glycerol, 0.01% Chaps) supplemented with 5 mM MgCl2. Chemical modification of GST-mCAD Modification of lysine residues of free GST-mCAD and the GST-mCAD/hICAD-L complex was performed using 20 and 60 µM, respectively, TNBS in 10 mM Tris–HCl pH 8.0, 100 mM NaCl, 1 mM EDTA, 1 mM DTT, 5% glycerol and 0.01% Chaps. For modification of the tyrosine residues 50 and 150 mM, respectively, NAI in 10 mM MES–NaOH pH 7.0 were used. To investigate protection against modification by TNBS or NAI, free GST-mCAD was also modified in the presence of 120 ng/µl assay solution of plasmid DNA (pBSK-VDEX). To analyse the residual activity of the modified nuclease, aliquots of the modification reaction mixtures were transferred into buffer C after defined time intervals, and incubated with caspase-3 (for reasons of standardisation always added) for 10 min at 37°C. In the case of modification of the GST-mCAD/hICAD-L complex and of free GST-mCAD in the absence of DNA, the aliquots were supplemented with plasmid pBSK-VDEX as substrate (25 ng/µl assay solution). To start the DNA cleavage reaction, MgCl2 was added to a final concentration of 5 mM and this reaction mix was then incubated for 10 min at 37°C. The cleavage products were analysed on a 0.8% TBE (100 mM Tris–HCl pH 8.3, 100 mM borate, 2.5 mM EDTA) agarose gel containing 0.05 µg/ml ethidium bromide. Site-directed mutagenesis Site-directed mutagenesis of GST-mCAD was performed as described by Kirsch and Joly (21). In brief, a first PCR was performed using a mutagenic primer and an appropriate reverse primer with pGEX-2T-mCAD as template and Pfu DNA polymerase. Then, a second PCR was performed using purified product from the first reaction as megaprimers for an inverse PCR following the instructions of the QuikChange protocol (Stratagene). In vitro GST-mCAD activity assays For the in vitro CAD activity assay, aliquots of dialysed free GST-mCAD were incubated in buffer C supplemented with 5 mM MgCl2 for defined time intervals at 37°C, using 25 ng/µl assay solution (10 nM final concentration) plasmid DNA (pBSK-VDEX). Cleavage products were analysed on a 0.8% TBE–agarose gel containing 0.05 µg/ml ethidium bromide. DNA–cellulose binding assay DNA binding of GST-mCAD variants was investigated by incubating caspase-3-treated GST-CAD/hICAD-L complex with 50 µl of a DNA–cellulose suspension in buffer B for 25 min at 4°C. Bound protein was washed once with 500 µl of buffer A supplemented with 5 mM DTT, 10% glycerol, 0.01% Triton X-100 and subsequently eluted by incubating the sample for 5 min at 95°C in 10 µl of SDS-gel loading buffer (160 mM Tris–HCl pH 6.8, 2% SDS, 5% β-mercaptoethanol, 40% glycerol and 0.1% bromophenol blue) and analysed by SDS–PAGE. Size-exclusion chromatography with GST-mCAD variants Size-exclusion chromatography with GST-mCAD and its variants was performed on a Superose 12 gel filtration column (24 ml bed volume, Pharmacia) equilibrated with 20 mM HEPES–NaOH pH 8.0, 150 mM KCl, 1 mM EDTA and 5 mM MgCl2 using a Merck-Hitachi HPLC system. Twenty micrograms of each variant were loaded onto the column and fractions of 1 ml were collected at a flow rate of 0.5 ml/min. Twenty-microlitre aliquots of the fractions were incubated in buffer C containing 5 mM MgCl2 for 1.75 h at 37°C with 20 ng/µl assay solution plasmid DNA (pBSK-VDEX). Cleavage products were analysed by agarose gel electrophoresis as described above. RESULTS Inactivation of CAD by chemical modification with TNBS and NAI and protection from chemical modification by DNA and ICAD-L Modification of free GST-mCAD with the lysine-specific reagent TNBS and the tyrosine-specific reagent NAI leads to an inhibition of the nucleolytic activity of the enzyme in a time- and dose-dependent manner, suggesting that one or more lysine and tyrosine residues are involved in the DNA cleavage reaction catalysed by CAD (Fig. 1). The presence of the inhibitor hICAD-L or of DNA during the TNBS- and NAI-modification reactions protects the nuclease from being modified, indicating that in the enzyme/substrate and enzyme/inhibitor complexes the essential lysine and tyrosine residues are in close proximity to both, the DNA substrate and the inhibitor, respectively, and therefore, not readily accessible for modification (data not shown). Effects of the substitution of conserved lysine and tyrosine residues on DNA cleavage by CAD An amino acid sequence alignment reveals that three out of a total of 19 lysine residues (Lys155, Lys301, Lys310) and two out of a total of 13 tyrosine residues (Tyr170, Tyr247) of murine CAD are fully conserved among the apoptotic nucleases from five different species and that all are located in the C-terminal catalytic domain of the enzyme (Fig. 2A). The evolutionary conservation of these residues and the results obtained in the chemical modification experiments prompted us to exchange them by site-directed mutagenesis to glutamine and phenylalanine, respectively. We have chosen these amino acid exchanges in order to test the importance of the functional group (a positive charge and a hydroxy group, respectively) and at the same time to avoid as much as possible any changes in protein integrity caused by an amino acid substitution that would alter the size and polarity of the side chain too much. DNA cleavage experiments with the mCAD variants produced in E.coli and purified to near homogeneity (Fig. 2B) demonstrate that all of the exchanged lysine and tyrosine residues are more or less important for DNA cleavage by CAD (Fig. 2C and Table 1), except for the Y170F variant, which retains wild-type activity. The strongest effect on activity is observed with the variant K155Q, which does not show any detectable DNA cleavage, even after prolonged incubation of the substrate DNA with high amounts of the enzyme. K301Q, K310Q and Y247F exhibit between 1 and 2% of wild-type mCAD activity as measured by the disappearance of supercoiled plasmid DNA. The variant Y170F retains wild-type activity with respect to its initial attack on supercoiled plasmid DNA, but displays a drastically altered cleavage pattern compared with wild-type mCAD by producing defined fragments rather than a ‘smear’. Effects of the substitution of conserved histidine residues on DNA cleavage by CAD In addition to the mutational analysis of conserved lysine and tyrosine residues of CAD, we have produced and analysed new CAD variants with substitutions of conserved histidine residues, which had been shown previously to be important for DNA cleavage (16,17). The residues His242, His263, His308 and His313 were substituted by Asp/Glu and Arg. With the exception of H313R, all variants showed little if any nucleolytic activity (Table 1). His313 of murine CAD, which is conserved in all known caspase−αχτιϖατεδ DNases from vertebrate organisms, has its counterpart in Arg418 of Drosophila CAD (Drep4) (17), which may explain why the substitution of His313 by Arg in murine CAD does not change the catalytic efficiency of the nuclease. However, like the variants Y170F (this work), H308N and H242N (17), the H313R variant produces a pattern of defined fragments when cleaving DNA (data not shown), pointing towards an involvement of His313 in DNA binding and/or processivity. CAD variant H308D exhibits residual DNA cleavage activity at low pH Among all variants with His→Asp/Glu substitutions, H308D is the only one with a residual cleavage activity at pH 5.0. Under these conditions all other variants show little if any activity, and the activity of the wild-type enzyme is much reduced compared with the activity at neutral pH (Fig. 3). At pH 5.0 the carboxy group of the aspartate residue is likely to be partially protonated (assuming a pKa ∼4.7), as is the histidine residue at pH 7.0 (assuming a pKa ∼6.4). Therefore, the protonated carboxy group could take over the putative function of the original histidine residue at this position in acting as the general acid in the mechanism of the CAD-catalysed DNA cleavage by protonating the leaving group after cleavage of a phosphodiester bond. Binding of CAD variants to DNA–cellulose In order to investigate DNA binding by the murine CAD variants, we have carried out a DNA–cellulose binding assay (16). To this end, GST-mCAD/hICAD-L mutant complexes were treated with caspase-3 and directly applied to the DNA–cellulose resin. Using this method, all GST-mCAD variants were shown to be able to bind to DNA–cellulose (Table 1), whereas the GST-mCAD/hICAD-L complex (data not shown) and GST itself, which were used as controls, did not bind to DNA (Fig. 4), demonstrating that all the GST-mCAD variants produced were not generally affected in DNA binding. CAD variants Y170F and H313R produce a defined DNA cleavage pattern As already observed for the substitution of His308 and His242 by Asn in mCAD (17), substitution of Tyr170 by Phe and His313 by Arg has a profound effect on the DNA cleavage mode compared with wild-type mCAD. The variants produce a well-defined cleavage pattern when degrading plasmid DNA under conditions where wild-type GST-mCAD produces randomly cut DNA fragments appearing as a ‘smear’. In order to exclude that this pattern was caused by trace amounts of an E.coli protein co-purified with the GST-mCAD variants and to confirm that the altered cleavage behaviour is an intrinsic property of the variant enzymes, we have expressed one variant (Y170F) also in 293-human embryonic kidney cells. The variant expressed in 293 cells, similar to the protein produced in E.coli, after purification produces a defined DNA cleavage pattern, whereas wild-type CAD produces randomly cut DNA fragments (data not shown). Since the activity of the Y170F and H313R variants in attacking supercoiled plasmid DNA is similar to that of wild-type CAD and since their binding to DNA is unaffected, it seems likely that Tyr170 and His313, together with His308 and probably His242, are important for the non-specific cleavage of substrate DNA by mCAD. Oligomerisation of GST-mCAD variants After dissociation from the complex with the inhibitor ICAD-L, CAD is known to oligomerise into its active form (7). In order to see if the GST-mCAD variants produced are still able to oligomerise, we have carried out gel filtration experiments with each variant using a Superose 12 column. Under the conditions applied, all variants displayed a similar elution volume (Table 1), with a peak corresponding to the void volume (2 MDa) of the column (data not shown), indicating that the drop in activity measured in the plasmid DNA cleavage assays is not due to impaired oligomerisation of the variant enzymes but rather a consequence of localised changes at or in close proximity to the active site of CAD. DISCUSSION The CAD/DFF40 plays a key role in DNA fragmentation during programmed cell death. Primary sequence analysis suggested that CAD belongs to a new nuclease family, since no sequence similarity with known nuclease families could be detected. To identify functionally important amino acid residues in CAD, DEPC-modification studies had been performed which strongly suggested that histidine residues might be involved in DNA binding and cleavage by this enzyme; not, however, in the interaction with ICAD-L (16,17). In a complementary mutational analysis, histidine residues were identified that might be essential for DNA cleavage by CAD (16,17). We have now extended these studies to find out whether other amino acid residues are essential for DNA cleavage by CAD, and have focused on lysine and tyrosine residues. In addition, we have produced new CAD variants to further analyse the role of histidine residues known to be involved in DNA cleavage by CAD. Our modification experiments demonstrate that lysine and tyrosine residues are required for DNA cleavage and that they are protected from modification by the presence of DNA or the inhibitor ICAD-L. It had been shown previously that the presence of DNA protected CAD from modification by DEPC, indicating that some of the histidine residues are in close proximity to the DNA substrate and inaccessible to the modifying compound. The inhibitory protein ICAD-L, in contrast, did not protect the nuclease from DEPC modification, leading to the conclusion that the substrate binding site and the inhibitor binding site of murine CAD are located on different regions (16). Our finding that TNBS and NAI modification of lysine and tyrosine residues is prevented by the presence of DNA as well as the inhibitory protein hICAD-L, suggests that at least one lysine and one tyrosine residue of CAD, in contrast to histidine residues, contribute directly or indirectly, not only to the substrate binding interface, but also to the inhibitor binding interface of murine CAD. It is known from previous structural and mutational analyses that conserved lysine residues (Lys12, Lys21, Lys35) in the N-terminal regulatory domain of CAD (CAD- or CIDE-N domain) contribute to the interaction with the N-terminal domain of ICAD. Single amino acid substitutions of these lysine residues by alanine had no effect on the nucleolytic activity and complex formation of CAD with ICAD-L; however, the double and triple mutants were proportionally more affected by the amino acid substitutions, with the latter being almost inactive and virtually unable to bind to ICAD-L (11,14). The successive loss of activity is a consequence of the inability of these CAD mutants to bind to the chaperone and inhibitor ICAD-L, which results in the formation of a catalytically incompetent nuclease, and not a consequence of the substitution of a critical active site residue. These results taken together suggest that the N-terminal domain of CAD and the region up to amino acid residue 241 harbours the binding site of CAD for ICAD-L (Fig. 5). In many different nucleases, lysine residues are part of the enzyme’s active site (22–31), where they can fulfil several functions. They can serve to correctly position a scissile phosphodiester bond and/or to stabilise the pentacovalent transition state during the enzymatic process, as well as to direct the attacking water molecule for an in-line attack on the phosphodiester bond. For example, in most type II restriction enzymes, a lysine residue is found in the catalytic PD....D/EXK-motif, where it functions mainly to position the attacking water molecule and to stabilise the transition state (reviewed in 32). In murine CAD we have identified by chemical modification, in conjunction with alignment-guided site-directed mutagenesis, three catalytically important lysine residues, Lys155, Lys301 and Lys310, that are located in the C-terminal catalytic domain of the enzyme and are fully conserved in all known CADs from five different species, namely mouse, rat, man, zebrafish and the fruitfly (Fig. 2). CAD variants with the lysine residues substituted by glutamine all exhibit a strongly reduced catalytic activity, with the variant K155Q exhibiting no detectable DNA cleavage activity, indicating that this residue might be particularly important for catalysis, possibly by transition state stabilisation. Tyrosine residues are also often part of the active site of a nuclease where they are involved in DNA binding by forming stacking interactions with a base or a sugar moiety of the nucleic acid. For example, Tyr76, a key DNA-binding residue of DNase I, which is part of a small loop that fills the minor groove of the DNA, forms a stacking interaction with a deoxyribose and thus plays an important role in the coupling of DNA recognition to phosphodiester bond cleavage by DNase I (33–36). In Serratia nuclease, Tyr76 can only be substituted by phenylalanine to preserve activity, presumably because it is involved in a stacking interaction with a base of the nucleic acid substrate (26,37,38). In the case of topoisomerases Ia and IIa the active site tyrosine residue attacks the scissile phosphodiester bond and forms a covalent intermediate in the mechanism of breakage and rejoining of the DNA (39). The tyrosine residues Tyr170 and Tyr247 of murine CAD when exchanged to phenylalanine show very different effects. The variant Y242F, similar to the variants K301Q and K310Q, shows a strongly reduced cleavage activity, which could mean that Tyr242, together with Lys301 and Lys310, contributes to productive substrate binding by correctly positioning the DNA in the substrate binding site (note that all variants studied here are not deficient in DNA binding as measured by the DNA–cellulose binding assay). The variants Y170F and H313R, in contrast, retain wild-type activity with respect to the cleavage of supercoiled plasmid DNA, but in the course of the cleavage reaction produce a pattern of fragments of defined length instead of randomly cut fragments as observed for wild-type murine CAD. It is likely from these results that Tyr170 and His313, probably together with His308 and His242 (17), for which a similar observation was made, contribute to a more non-specific DNA cleavage mode, which leads to a ‘smear’-like product distribution, but do not participate in catalysis per se. Murine CAD binds to DNA in the absence of Mg2+, which, however, is needed as the divalent cation cofactor for the DNA cleavage reaction. Using a DNA–cellulose binding assay, it could be demonstrated that the substitution of critical histidine residues abolishes DNA cleavage, not, however, DNA binding of CAD, except for the variant H242A (16). In order to analyse DNA binding by CAD variants with exchanged lysine and tyrosine residues, we have conducted a DNA–cellulose-binding assay with these variants. All of them bind to DNA–cellulose with the same efficiency (as far as this can be measured with such a semi-quantitative assay), indicating that the mutations introduced do not affect the overall DNA-binding capability of CAD. However, this does not exclude that the conserved lysine and tyrosine residues under investigation each contribute to substrate binding by CAD and that their individual substitution leads to a diminished nucleolytic activity. In nucleases, several residues are involved in DNA binding, often by formation of electrostatic interactions between positively charged side chains and the negatively charged phosphodiester backbone, or stacking interactions between aromatic side chains and the bases or the sugar of the nucleic acid substrate. The substitution of one of these residues can lead to a decrease in the catalytic efficiency of the nuclease, but does not necessarily prevent DNA binding. Instead, it could be that in these variants the substrate is not properly positioned, due to the absence of a contact required for a productive interaction. CONCLUSION Our investigation of the role of conserved lysine and tyrosine residues of the CAD demonstrates that several lysine and tyrosine residues contribute to DNA binding and cleavage by CAD, and to the inhibition of CAD activity by binding to the inhibitor ICAD-L. Lys155, when substituted by glutamine, shows the most dramatic decrease in nucleolytic activity, emphasising its particular role in the mechanism of DNA cleavage, which could be positioning of the attacking nucleophile or transition state stabilisation. Lys301, Lys310 and Tyr247 also seem to be part of the enzyme/DNA interface, where they could be involved in binding and positioning of the substrate. Tyr170 and His313, like His308 and His242, seem to be involved in DNA binding in such a way as to guarantee a non-specific DNA cleavage mode, which may be associated with the processivity of the enzyme. Lysine and tyrosine residues co-operate with histidine residues, which we had begun to analyse before (17), in DNA cleavage. Because of the large effects observed with substitution of His263 by asparagine, we had suggested that this histidine residue might act as a general base in the mechanism of DNA cleavage by CAD. We now have evidence that His308 could be the general acid. It is noteworthy that all residues which were shown to affect the DNA cleavage activity when exchanged are located between Lys155 and His313 (Fig. 5), which, therefore, allows one to provisionally define the boundary of the catalytic domain. The function of the essential Mg2+ has not yet been addressed. For this purpose it will be important to identify the amino acid residues involved in Mg2+ binding, which is underway in our laboratory. ACKNOWLEDGEMENTS We thank Ms Ute Konradi for expert technical assistance. This work has been supported by a grant from the Deutsche Forschungsgemeinschaft (Pi 122/16-1) and the Fonds der Chemischen Industrie. S.R.S. is a member of the Graduiertenkolleg ‘Biochemie von Nukleoproteinkomplexen’. O.G.’s stay at Giessen was supported by the Deutsche Akademische Austauschdienst. * To whom correspondence should be addressed. Tel: +49 641 99 35404; Fax: +49 641 99 35409; Email: [email protected] View largeDownload slide Figure 1. Chemical modification of murine CAD by TNBS and NAI. Free GST-mCAD was modified by TNBS (top) and NAI (bottom) for the indicated time (1.5, 3, 6, 7.5 and 9 min). Subsequently the residual nucleolytic activities of the aliquots withdrawn from the modification reaction mixtures at the indicated time points were measured by a plasmid DNA cleavage assay (10 min). TNBS and NAI inactivate free GST-mCAD in a time- and dose-dependent manner. (ST, length standard; S, substrate DNA; C, control reaction; oc, open circular DNA; lin, linear DNA; sc, supercoiled DNA.) View largeDownload slide Figure 1. Chemical modification of murine CAD by TNBS and NAI. Free GST-mCAD was modified by TNBS (top) and NAI (bottom) for the indicated time (1.5, 3, 6, 7.5 and 9 min). Subsequently the residual nucleolytic activities of the aliquots withdrawn from the modification reaction mixtures at the indicated time points were measured by a plasmid DNA cleavage assay (10 min). TNBS and NAI inactivate free GST-mCAD in a time- and dose-dependent manner. (ST, length standard; S, substrate DNA; C, control reaction; oc, open circular DNA; lin, linear DNA; sc, supercoiled DNA.) View largeDownload slide Figure 2. Mutational analysis of variants with exchanged conserved lysine and tyrosine residues of GST-mCAD. (A) Amino acid residues Lys155, Lys301 and Lys310 as well as Tyr170 and Tyr247, which were exchanged to glutamine or phenylalanine, respectively, are fully conserved among the apoptotic nucleases known from the five indicated species [Mus musculus (GenBank accession nos AB009377, NM_007859), Rattus norvegicus (GenBank accession no. AF136598), Homo sapiens (GenBank accession nos AF064019, AF039210, AB013918, NM_004402), Danio rerio (GenBank accession no. AF286179) and Drosophila melanogaster (GenBank accession nos AF149797, AB036773)]. Shading is according to the Blosum 62 scoring matrix, with black shading for 100%, dark grey for 80% and light grey for 60% conserved amino acid residues. (B) SDS–PAGE analysis of variants of GST-mCAD that were produced as a complex with hICAD-L in E.coli and activated by treating the complex with recombinant caspase-3. (C) DNA cleavage activity of the GST-mCAD variants were measured by the disappearance of supercoiled plasmid DNA, analysed by agarose gel electrophoresis. GST-mCAD-Y170F is the only variant that retains wild-type activity with respect to the cleavage of supercoiled DNA, whereas all other variants exhibit strongly reduced cleavage activities. GST-mCAD-Y170F produces a pattern of fragments of defined length instead of randomly cut fragments appearing as a ‘smear’ (see wild-type GST-mCAD). Note the different concentrations of the variants as indicated. View largeDownload slide Figure 2. Mutational analysis of variants with exchanged conserved lysine and tyrosine residues of GST-mCAD. (A) Amino acid residues Lys155, Lys301 and Lys310 as well as Tyr170 and Tyr247, which were exchanged to glutamine or phenylalanine, respectively, are fully conserved among the apoptotic nucleases known from the five indicated species [Mus musculus (GenBank accession nos AB009377, NM_007859), Rattus norvegicus (GenBank accession no. AF136598), Homo sapiens (GenBank accession nos AF064019, AF039210, AB013918, NM_004402), Danio rerio (GenBank accession no. AF286179) and Drosophila melanogaster (GenBank accession nos AF149797, AB036773)]. Shading is according to the Blosum 62 scoring matrix, with black shading for 100%, dark grey for 80% and light grey for 60% conserved amino acid residues. (B) SDS–PAGE analysis of variants of GST-mCAD that were produced as a complex with hICAD-L in E.coli and activated by treating the complex with recombinant caspase-3. (C) DNA cleavage activity of the GST-mCAD variants were measured by the disappearance of supercoiled plasmid DNA, analysed by agarose gel electrophoresis. GST-mCAD-Y170F is the only variant that retains wild-type activity with respect to the cleavage of supercoiled DNA, whereas all other variants exhibit strongly reduced cleavage activities. GST-mCAD-Y170F produces a pattern of fragments of defined length instead of randomly cut fragments appearing as a ‘smear’ (see wild-type GST-mCAD). Note the different concentrations of the variants as indicated. View largeDownload slide Figure 3. Residual DNA cleavage activities of variants of GST-mCAD with His→Asp/Glu substitutions. Variant H308D is the only variant with a measurable activity at pH 5.0. All other variants, except wild-type GST-mCAD, are virtually inactive at this low pH. View largeDownload slide Figure 3. Residual DNA cleavage activities of variants of GST-mCAD with His→Asp/Glu substitutions. Variant H308D is the only variant with a measurable activity at pH 5.0. All other variants, except wild-type GST-mCAD, are virtually inactive at this low pH. View largeDownload slide Figure 4. DNA binding by the GST-mCAD variants. DNA binding of the GST-mCAD variants was investigated using a DNA–cellulose binding assay (16). Caspase-3-treated GST-CAD/hICAD-L complex (2.5 µg) and GST (2.5 µg) were incubated with 50 µl of a DNA–cellulose suspension. After washing, bound proteins were eluted by incubating the suspension for 5 min at 95°C in 10 µl SDS-gel loading buffer and subsequently analysed by SDS–PAGE. As can be seen, all GST-mCAD variants bind to DNA–cellulose, whereas GST alone does not. View largeDownload slide Figure 4. DNA binding by the GST-mCAD variants. DNA binding of the GST-mCAD variants was investigated using a DNA–cellulose binding assay (16). Caspase-3-treated GST-CAD/hICAD-L complex (2.5 µg) and GST (2.5 µg) were incubated with 50 µl of a DNA–cellulose suspension. After washing, bound proteins were eluted by incubating the suspension for 5 min at 95°C in 10 µl SDS-gel loading buffer and subsequently analysed by SDS–PAGE. As can be seen, all GST-mCAD variants bind to DNA–cellulose, whereas GST alone does not. View largeDownload slide Figure 5. Variants of CAD with reduced catalytic activity. (A) Inohara et al. (11) and Otomo et al. (14) have identified three conserved lysine residues (Lys12, Lys21 and Lys35) in the N-terminal domain of CAD (amino acids 1–83) that contribute to ICAD-L binding. Inohara et al. (11) have also identified three conserved residues (Gly55, Phe63 and Trp81) in the N-terminal domain of CAD that seem to enhance the nuclease activity of CAD without being essential for catalysis. According to their results, the catalytic nuclease domain is located in the C-terminal region of the protein (amino acids 84–344), since the Δ1–83 variant of CAD exhibits a moderate nucleolytic activity, whereas mutant forms with deletions in the C-terminal domain (Δ290–344, Δ162–344 and Δ84–344) have no nuclease activity. It must be emphasised that Otomo et al. (14) have reported contradictory results, according to which the C-terminal domain of CAD (Δ1–83) does not show any nucleolytic activity. (B) The C-terminal domain of CAD contains several histidine, lysine and tyrosine residues, which have been identified as being catalytically relevant by chemical modification and alignment-guided site-directed mutagenesis (16,17; this work). Variants of CAD with substitutions of the amino acid residues marked with an asterisk (Tyr170, His242, His308 and His313) are active but produce a defined cleavage pattern when hydrolysing DNA. (C) It can be concluded from the structural studies as well as the chemical modification and mutational analyses that the region of CAD interacting with ICAD-L, besides the known interaction of the N-terminal domain (12–15), includes the amino acid residues of CAD up to residue 241, whereas the DNA binding site is between amino acid residues Lys155 and His313. View largeDownload slide Figure 5. Variants of CAD with reduced catalytic activity. (A) Inohara et al. (11) and Otomo et al. (14) have identified three conserved lysine residues (Lys12, Lys21 and Lys35) in the N-terminal domain of CAD (amino acids 1–83) that contribute to ICAD-L binding. Inohara et al. (11) have also identified three conserved residues (Gly55, Phe63 and Trp81) in the N-terminal domain of CAD that seem to enhance the nuclease activity of CAD without being essential for catalysis. According to their results, the catalytic nuclease domain is located in the C-terminal region of the protein (amino acids 84–344), since the Δ1–83 variant of CAD exhibits a moderate nucleolytic activity, whereas mutant forms with deletions in the C-terminal domain (Δ290–344, Δ162–344 and Δ84–344) have no nuclease activity. It must be emphasised that Otomo et al. (14) have reported contradictory results, according to which the C-terminal domain of CAD (Δ1–83) does not show any nucleolytic activity. (B) The C-terminal domain of CAD contains several histidine, lysine and tyrosine residues, which have been identified as being catalytically relevant by chemical modification and alignment-guided site-directed mutagenesis (16,17; this work). Variants of CAD with substitutions of the amino acid residues marked with an asterisk (Tyr170, His242, His308 and His313) are active but produce a defined cleavage pattern when hydrolysing DNA. (C) It can be concluded from the structural studies as well as the chemical modification and mutational analyses that the region of CAD interacting with ICAD-L, besides the known interaction of the N-terminal domain (12–15), includes the amino acid residues of CAD up to residue 241, whereas the DNA binding site is between amino acid residues Lys155 and His313. Table 1. Properties of variants with substitution of conserved histidine, lysine and tyrosine residues in the C-terminal catalytic domain of the murine CAD Residue  Substitution  ICAD-L bindinga  Relative DNA cleavage activity (%)b  Oligomeric state  DNA bindingc  Remark  Wild-type  –  +  100  wt  wt    Lys155  Gln  +  n.d.c.  wt  wt    Lys301  Gln  +  1.4  wt  wt    Lys310  Gln  +  2.3  wt  wt    Tyr170  Phe  +  107.0  wt  wt  Fragment pattern  Tyr247  Phe  +  1.2  wt  wt    His127  Asnd  +  25.8  wt  n.d.e    His242  Asnd  +  6.3  Deviating  n.d.e  Fragment pattern    Glu  +  <1.0  wt  wt      Arg  +  <1.0  wt  wt    His263  Asnd  +  0.7  wt  n.d.e      Asp  +  <1.0  wt  wt      Arg  +  <1.0  wt  wt    His304  Asnd  +  14.5  wt  n.d.e    His308  Asnd  +  9.3  wt  n.d.e  Fragment pattern    Asp  +  ∼1–2  wt  wt  Residual activity at pH 5.0    Arg  +  <1.0  wt  wt    His313  Asnd  +  1.3  wt  n.d.e      Asp  +  <1.0  wt  wt      Arg  +  109.0  wt  wt  Fragment pattern  Residue  Substitution  ICAD-L bindinga  Relative DNA cleavage activity (%)b  Oligomeric state  DNA bindingc  Remark  Wild-type  –  +  100  wt  wt    Lys155  Gln  +  n.d.c.  wt  wt    Lys301  Gln  +  1.4  wt  wt    Lys310  Gln  +  2.3  wt  wt    Tyr170  Phe  +  107.0  wt  wt  Fragment pattern  Tyr247  Phe  +  1.2  wt  wt    His127  Asnd  +  25.8  wt  n.d.e    His242  Asnd  +  6.3  Deviating  n.d.e  Fragment pattern    Glu  +  <1.0  wt  wt      Arg  +  <1.0  wt  wt    His263  Asnd  +  0.7  wt  n.d.e      Asp  +  <1.0  wt  wt      Arg  +  <1.0  wt  wt    His304  Asnd  +  14.5  wt  n.d.e    His308  Asnd  +  9.3  wt  n.d.e  Fragment pattern    Asp  +  ∼1–2  wt  wt  Residual activity at pH 5.0    Arg  +  <1.0  wt  wt    His313  Asnd  +  1.3  wt  n.d.e      Asp  +  <1.0  wt  wt      Arg  +  109.0  wt  wt  Fragment pattern  n.d., not determined; n.d.c., no detectable cleavage; wt, wild-type-like. aICAD-L binding measured by co-purification of untagged hICAD-L with GST-mCAD. bMeasured by the disappearance of supercoiled plasmid DNA in steady state cleavage experiments. cBinding of free GST-mCAD to DNA–cellulose. dThe properties of these variants were described in Meiss et al. 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