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Single molecule enzymology: watching the reaction
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Single molecule enzymology: watching the reaction Anne Gershenson1 Single molecule optical microscopy can directly monitor substrate turnover by individual enzymes revealing the underlying distribution of reaction rates and enzyme conformations. These techniques are particularly useful for assessing cooperativity in multi-subunit enzymes such as b-galactosidase, and for directly monitoring how ligand and substrate binding alter dynamic equilibria. Recent investigations of HIV reverse transcriptase have reiterated the importance of single molecule microscopy for determining how proteins move on oligonucleotides and how ligands and inhibitors affect motion. Similar investigations of membrane active enzymes allow direct imaging of protein–membrane interactions. For a large variety of systems, single molecule enzymology provides unprecedented images of how enzymes interact with their substrates and the differences between individual enzymes in a population. Address Chemistry Department, Brandeis University, MS 015, 415 South Street, Waltham, MA 02454, USA Corresponding author: Gershenson, Anne ([email protected]) 1 Current address: Department of Biochemistry & Molecular Biology, 1230 Lederle Graduate Research Tower, University of Massachusetts, Amherst, 710 N. Pleasant Street, Amherst, MA 01003, USA.
Current Opinion in Chemical Biology 2009, 13:436–442 This review comes from a themed issue on Mechanisms Edited by Catherine L. Drennan and Joseph T. Jarrett Available online 24th July 2009 1367-5931/$ – see front matter # 2009 Elsevier Ltd. All rights reserved. DOI 10.1016/j.cbpa.2009.06.011
Introduction Over the past 20 years, single molecule optical and force methods have revealed how proteins move on microtubules and oligonucleotides, and have helped connect protein motion to chemical reactions such as the hydrolysis of ATP. The ability to visualize substrate turnover by individual enzymes has also confirmed that many enzymes are conformationally heterogeneous displaying either static heterogeneity where individual enzyme molecules exhibit different catalytic rates, or dynamic heterogeneity, where the catalytic rate of an individual enzyme changes over time. In order to determine the distribution of states within a population, single molecule experiments require data from hundreds of enzyme turnovers. Collecting enough Current Opinion in Chemical Biology 2009, 13:436–442
data for good statistics can be tedious and is facilitated by methods for interrogating large numbers of single molecules at the same time, that is, single molecule multiplexing. Traditionally, multiplexing has been achieved by immobilizing enzymes or substrates (e.g. DNA) on a surface and using conventional and/or fluorescence microscopy to image the molecules (see [1–4] for recent reviews of single molecule optical and force techniques). Recently, a number of new methods for multiplexing have been developed and these are described in Box 1. Several of these methods avoid direct enzyme or substrate immobilization and are likely to increase the number of systems amenable to single molecule techniques. Recent reviews have covered the extensive single molecule literature on systems such as polymerases, ribosomes, helicases and motor proteins [5–8]. This review focuses on recent work on soluble enzymes with small, soluble substrate; highlights important new single molecule work on HIV reverse transcriptase and pays particular attention to the emerging single molecule literature on how peripheral membrane proteins interact with lipid bilayers.
Soluble enzyme cooperativity and heterogeneity b-galactosidase, Inhibition & Enzyme Heterogeneity
A relevant question for oligomeric enzymes is the role of allostery in enzyme activity and inhibition. Escherichia coli b-galactosidase is only active as a tetramer, and the four active sites turnover substrates independently [9,10,11]. One might, therefore, expect that in the presence of inhibitors single b-galactosidase molecules would have 4 distinct activity levels corresponding to 0, 1, 2 or 3 bound inhibitors. However, individual b-galactosidase enzymes exhibited cooperative inhibitor binding and release, switching between states with 0 or 4 bound inhibitors [10]. One surprise was that the activity of single b-galactosidases often significantly changed after inhibitor release, a sign of dynamic heterogeneity caused by protein conformational changes [10]. b-galactosidase dynamic heterogeneity was previously quantified in experiments where individual enzymes immobilized on beads exhibited both long and short waiting times between substrate turnovers [9]. By contrast, experiments in microchambers (Box 1) show that while b-galactosidase molecules display a large distribution of turnover velocities, the velocity of a single b-galactosidase is stable for hundreds of seconds indicating static heterogeneity [12]. Why are different types of heterogeneity observed? Single turnovers were detected in the bead-based experiments, while in the microchambers the results of www.sciencedirect.com
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multiple turnovers were recorded every 15 s. This suggests that on short timescales at ambient temperatures the conformation of a single b-galactosidase enzyme fluctuates around an average conformation with a relatively well-defined long term catalytic rate. This long term rate can change in response to large perturbations such as inhibitor binding. Adenylate kinase enzyme activity and dynamic heterogeneity. All of the evidence for b-galactosidase conformational heterogeneity is indirect because the fluorescent label is on the substrate. Single pair Fo¨rster resonance energy transfer (spFRET) can be used to directly monitor large, functionally important conformational changes and to assess enzyme conformational heterogeneity. In these experiments, a protein is labeled with both a donor and an acceptor fluorophore and changes in the distance between the two probes alters the fluorescence signal (for a guide to spFRET see [13]). Two recent studies of adenylate kinase (AK) used spFRET to measure AK conformational distributions and time-resolved conformational changes in the presence and absence of substrate analogues [14,15]. AK catalyzes the reversible conversion of ATP/AMP to two ADP molecules helping to maintain the energy balance in cells. The AK active site has two flexible lids, the ATP lid in one domain and the AMP lid in the second domain [15]. SpFRET as well as X-ray crystal structures reveal that apo-AK can sample a large distribution of lid conformations, from fully opened to fully closed [14,15]. In the presence of substrate analogue or inhibitor the spFRET distributions shifted to favour the closed conformation [14,15]. For E. coli AK this shift is caused by a doubling of the closing rate constant biasing the conformational distribution towards lid closure in the presence of AMP-PNP, substrate analogue and AMP, substrate [14], thus, demonstrating the utility of spFRET for
directly monitoring dynamic equilibria and the shifting of equilibria by substrates or ligands. AAA+-ATPase Mechanisms Members of the AAA+ ATPase superfamily are generally ring-shaped multisubunit complexes many of which couple ATP binding and hydrolysis to mechanical work. Systems studied using single molecule techniques include F1-ATPase [16,17,18,19,20,21,22], proteins involved in homologous recombination such as helicases [23], RecA [23– 25] and Rad51 [23,26] and the Clp family of unfoldases [27,28]. Recent, elegant single molecule experiments on F1-ATPase, the soluble rotary motor from the F1F0-ATP synthase that can either hydrolyze or synthesize ATP depending on the direction of rotation [29,30], have filled in some of the missing steps in the F1-ATPase mechanism. The minimal F1-ATPase functional complex consists of a g subunit, the rotor, inserted into the centre of the pseudohexameric a3b3 cylindrical stator composed of alternating a and b subunits. Attachment of a filament or beads to the g subunit allows rotation of individual F1ATPase complexes to be observed using conventional or fluorescence microscopy. Such experiments revealed that rotation of the rotor occurs in 1208 steps [31], and these steps have since been shown to consist of 808 and 408 substeps [16,32] (Figure 1). In general, one of the b subunits is empty, bE, one has ATP bound, bATP, and the third has ADP or ADP-Pi bound, bADP [33,34]. ATP binding to bE leads to ADP release from bADP, and the 808 substep during which ATP in the third subunit is hydrolyzed to ADP-Pi [16]. The temperature dependence of F1-ATPase stepping recently revealed that ADP release is rate limiting [17,22]. What about the 408 substep? Excess Pi in solution can drive 408 rotations in the opposite direction showing that Pi release drives the 408 substep [16]. In addition, fluorescently labeled
Box 1 Immobilizing or encapsulating single molecules To ensure that an individual molecule stays in the observation volume, time resolved single molecule microscopy requires limiting the volume available to each molecule. This has often been accomplished using immobilization. However, concerns about the effects of surfaces on enzyme activity as well as the difficulties inherent in selective immobilization have led to the development of encapsulation strategies. Immobilization or encapsulation also allows single molecules to be arrayed so that data from multiple molecules may be collected simultaneously helping to ensure good experimental statistics. A number of immobilization and encapsulation strategies are illustrated. (a) Single molecule experiments often use immobilized proteins (top left) or substrates (top right). (b) Immobilization of large numbers of DNA strands near a nanoscale barrier can be achieved by combining lipid attached DNA molecules (green on orange circles), nanobarriers and flow [26,49]. (c) In this scheme, the lipids diffuse towards the barrier and flow stretches the DNA creating a ‘DNA curtain’. Figure adapted from Fazio et al. [49]. Alternatively, enzymes may be encapsulated in microchambers at the ends of optical fibres, www.sciencedirect.com
where the illustrated reaction is for b-galactosidase hydrolyzing resorfin-b-galactopyranoside. There are hundreds of fibres in the bundles allowing multiplexing, but buffer exchange is complicated. This figure was adapted from Gorris et al. [10]. (d) Enzymes and large substrates such as DNA may be encapsulated in vesicles made porous by incorporating protein pores (left) or bad lipid packing (right). The vesicles are easily immobilized, and the holes in the membrane allow buffer exchange. This figure was adapted from Cisse et al. [24]. (e) Enzymes may be encapsulated in naturally porous virus capsids that can then be immobilized. E is the encapsulated enzyme, S and P, the substrate and product, respectively, can diffuse in and out of the capsid. This figure was adapted from Comellas-Aragones et al. [50]. (f) Enzymes may be encapsulated in aqueous droplets and, as shown in the time series from left to right, the droplets can be fused allowing buffer exchange [51]. The scale bar in the first frame is 1 mm (picture courtesy of LS Goldner). Droplets can also be multiplexed using a number of microfluidic architectures. Current Opinion in Chemical Biology 2009, 13:436–442
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ATP bound at 08 is released as ADP following a 2408 rotation (two 1208 steps). How is the energy gained from the chemical steps converted to mechanical work? Single molecule fluorescence polarization experiments have correlated nucleotide binding states with opening and closing of labeled b subunits suggesting that bending motions of the b subunits drive rotation [20] (Figure 1). It has also been suggested that interactions between the g rotor and both the bottom and top of the a3b3 stator drive rotations [18,19,35]. However, truncated g rotors that only sit on the top of the stator can still generate torque and processively rotate in the correct direction suggesting that biased diffusion may be important for F1-ATPase rotary Current Opinion in Chemical Biology 2009, 13:436–442
motion [18,19]. While more experiments on the details of the mechanical couplings remain to be done, single molecule F1-ATPase experiments are close to providing a complete, detailed molecular mechanism for the conversion of energy from chemical reactions to rotary motion.
Motion on a track and HIV reverse transcriptase (RT) To catalyze reactions on DNA or RNA enzymes must partition from solution unto quasi-one dimensional oligonucleotides and find their targets. Single molecule studies using fluorescently labeled, surface immobilized oligonucleotides and fluorescently labeled proteins have been particularly valuable for elucidating how enzymes bind to, travel on and manipulate oligonucleotides www.sciencedirect.com
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Figure 1
The proposed mechanism for F1-ATPase rotation. ATP binding to the open, empty b subunit, b1, triggers closing of b1. This closing leads to ADP release from the b2 subunit, located 1208 to the left of b1, and an 808 counter-clockwise substep by the g subunit (yellow teardrop). Subsequently inorganic phosphate, Pi, is released from b2 triggering the 408 substep and ATP is hydrolyzed by b3 allowing b3 to partially open. It is currently unclear whether hydrolysis of ATP and the associated b subunit conformational changes occur before or after the rotational substeps. Open, partially closed and fully closed b subunit conformations are shown schematically in red, green and blue, respectively. T indicates bound ATP and D indicates bound ADP. This figure is adapted from Masaike et al. [20].
[5,23,36]. In the past year, spFRET studies by Zhuang and co-workers have provided important insights into how labeled RT interacts with labeled oligonucleotides and how inhibition by noncompetitive non-nucleoside RT inhibitors alters these interactions [37,38]. Heterodimeric RT, which has both DNA polymerase and RNAse H activity, transcribes the viral genome from single stranded RNA to double stranded DNA allowing infected cells to replicate the virus [39]. Intriguingly, the distribution of RT locations on primer-template oligonucleotides as well as RT motion depend on the composition of the oligonucleotide hybrids [37,38]. On mimics for plus strand elongation RT predominantly bound in a polymerase competent manner with the fingers domain at the 30 end of the primer strand [37] (Figure 2). By contrast, RT predominantly bound in the opposite conformation, with the RNase H domain at the 30 end of the primer, for mimics of RNase H substrates. Experiments on hybrid primer strands composed of both RNA and DNA, demonstrate that the 50 composition of the primer strand determines the binding orientation. Thus, the binding orientation and position of RT help determine its function. How does RT find the primer ends and how does it assume the correct orientation? Time-resolved spFRET studies of RT–oligonucleotide interactions show that RT motion is rarely unidirectional and the protein slides back and forth along the track [38]. Sliding is integral to the RT mechanism and elongating RT slides back and forth even when it must displace an RNA strand. Nucelotides and inhibitors alter the dynamic equilibrium for sliding with the rate constant for sliding to the 50 end of the primer (kback) decreasing in the presence of cognate dNTPs and increasing in the presence of the non-nucleoside RT inhibitor nevirapine. In addition to sliding, RT can flip around the substrate to assume the correct orientation, for www.sciencedirect.com
example, flipping from an RNase H competent conformation with the from RNase H domain at the 30 end of the primer to a polymerase competent conformation with the fingers domain at the 30 end of the primer, while still remaining attached to the substrate [38]. Thus, these elegant experiments reveal the dynamic equilibria that are integral to RT function and how biasing sliding favours elongation or inhibition.
Peripheral membrane proteins and interfacial catalysis Substrate turnover by peripheral membrane proteins depends not only on the details of the chemical mechanism but also on how the enzymes partition to and interact with two-dimensional membranes. Single molecule fluorescence microscopy can directly monitor protein motions at the interface and a related techniques fluorescence correlation spectroscopy (FCS) can monitor the diffusion time scale that changes dramatically when small proteins bind much larger vesicles [40,41–43]. Membrane active enzymes can exhibit ‘substrate dilution inhibition’ where a sharp decrease in enzyme activity is observed as the substrate is diluted in the membrane. For extracellular Bacillus thuringiensis phosphatidylinositol specific phospholipase C (PI-PLC), which catalyzes cleavage of the sn-3-phosphodiester bond in phosphatidylinositol (PI), inhibition can occur even when 40% of the membrane is substrate suggesting the involvement of factors other than substrate scarcity [40]. FCS experiments on PI-PLC reveal that the affinity of B. thuringiensis PI-PLC for lipid vesicles initially parallels enzyme activity and Kd drops an order of magnitude for vesicles containing anionic lipids and 10% phosphatidylcholine (PC), an effector lipid that enhances activity [40]. However, unlike the activity, which is stable up to 50% PC and then precipitously drops, Kd continues to decrease as the Current Opinion in Chemical Biology 2009, 13:436–442
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Figure 2
Experimental setup and conformational distributions for RT spFRET experiments. (a) The substrate consisting of an acceptor labeled template strand and a primer strand were immobilized on a quartz coverslip and donor labeled RT was added. The donor and acceptor fluorescence were excited using total internal reflection fluorescence (TIRF) microscopy, which preferentially excites fluorophores near the surface. RT binding to the substrate leads to spFRET and acceptor fluorescence. H indicates the RT RNase H domain and F indicates the fingers domain. The 50 ends of the oligonucleotides are indicated by circles and the 30 ends are indicated by arrows. (b) When binding to plus strand synthesis mimics, DNA(primer)DNA(template), RT primarily binds in a polymerase competent orientation with the fingers domain near the 30 end of the primer resulting in high spFRET efficiencies when the donor is in the RNase H domain (top). RT binds RNA(primer)-DNA(template) substrates in the opposite orientation with the RNase H domain at the 30 end of the primer and low spFRET efficiency (bottom). DNA is shown in black, RNA is shown in red and the donor and acceptor fluorophores are shown in green and red, respectively. Figures are adapted from Abbondanzieri et al. [37].
PC content is increased to 90% indicating tight membrane binding. Parallel NMR studies of the phospholipid headgroups reveal that tight PI-PLC binding preferentially restricts the mobility of the PC headgroup [40]. Thus, substrate dilution inhibition likely arises from substrate scarcity as well as restrictions on protein and lipid mobility. Interactions of fluorescently labeled proteins with supported lipid bilayers and multilayers can be directly imaged with single molecule fluorescence microcopy [44,45,46]. Studies by Yang and co-workers on cobra phospholipase A2 (PLA2), a Ca2+-dependent phospholipase that catalyzes hydrolysis of the sn-2 ester of glycerophospholipids, recently demonstrated that binding and activity preferentially occur near membrane defects [44]. Similar experiments performed by Uji-i and co-workers on a mutant phospholipase A1 (PLA1) that catalyzes hydrolysis of the sn-1 ester of phospholipids also showed preferential binding and activity at defects [45]. Why this preference for defects? High membrane curvature associated with defects or lipid tubes on the top of multilayers probably increase substrate accessibility reducing the energy needed to pull the phospholipid headgroup into the active site [44,45]. PLA1 also showed a number of different interactions with the membranes ranging from fast, transient association to slower diffusion on top of multilayers to occasional immoCurrent Opinion in Chemical Biology 2009, 13:436–442
bilization. The diffusion of single active PLA1 molecules changed over time, transitioning from slow diffusion near membrane edges, probably associated with catalysis, to faster diffusion presumably associated with the search for a new substrate lipid [45]. Inactive PLA1 diffusion was less heterogeneous, generally displaying a diffusion coefficient similar to that expected for a lipid [45]. Similar lipid assisted scooting, where protein motion results from diffusion of a bound lipid, seems to be a general feature of protein–lipid interactions and has been observed in single molecule fluorescence studies of acyl-CoA binding protein bound to POPC giant unilamellar vesicles [47] and for the GRP1 pleckstrin homology domain on supported lipid bilayers [48]. Single molecule experiments have thus begun to correlate different modes of protein–membrane interactions with the catalytic cycle and to visualize binding hotspots associated with enzyme activity.
Conclusions Recent single molecule enzyme studies demonstrate the importance of these techniques for probing catalytically relevant conformational changes, the cooperativity of activity and inhibitor binding/release in oligomeric proteins as well as probing interaction between proteins and oligonucleotides or membranes. As highlighted by the work on b-galactosidase, RT and PI-PLC, single fluorescence molecule techniques are particularly useful for determining the molecular mechanisms of inhibition www.sciencedirect.com
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in complicated systems. The advent of new methods for protein encapsulation as well as microfluidic methods may obviate the need for surface immobilization, expanding the applicability of these methods.
Acknowledgements Research in the author’s laboratory is funded in part by a grant from the Alpha-1 Foundation and supported by a charitable donation from Talecris Biotherapeutics, Center for Science and Education.
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Single molecule enzymology: watching the reaction Anne Gershenson1 Single molecule optical microscopy can directly monitor substrate turnover by individual enzymes revealing the underlying distribution of reaction rates and enzyme conformations. These techniques are particularly useful for assessing cooperativity in multi-subunit enzymes such as b-galactosidase, and for directly monitoring how ligand and substrate binding alter dynamic equilibria. Recent investigations of HIV reverse transcriptase have reiterated the importance of single molecule microscopy for determining how proteins move on oligonucleotides and how ligands and inhibitors affect motion. Similar investigations of membrane active enzymes allow direct imaging of protein–membrane interactions. For a large variety of systems, single molecule enzymology provides unprecedented images of how enzymes interact with their substrates and the differences between individual enzymes in a population. Address Chemistry Department, Brandeis University, MS 015, 415 South Street, Waltham, MA 02454, USA Corresponding author: Gershenson, Anne ([email protected]) 1 Current address: Department of Biochemistry & Molecular Biology, 1230 Lederle Graduate Research Tower, University of Massachusetts, Amherst, 710 N. Pleasant Street, Amherst, MA 01003, USA.
Current Opinion in Chemical Biology 2009, 13:436–442 This review comes from a themed issue on Mechanisms Edited by Catherine L. Drennan and Joseph T. Jarrett Available online 24th July 2009 1367-5931/$ – see front matter # 2009 Elsevier Ltd. All rights reserved. DOI 10.1016/j.cbpa.2009.06.011
Introduction Over the past 20 years, single molecule optical and force methods have revealed how proteins move on microtubules and oligonucleotides, and have helped connect protein motion to chemical reactions such as the hydrolysis of ATP. The ability to visualize substrate turnover by individual enzymes has also confirmed that many enzymes are conformationally heterogeneous displaying either static heterogeneity where individual enzyme molecules exhibit different catalytic rates, or dynamic heterogeneity, where the catalytic rate of an individual enzyme changes over time. In order to determine the distribution of states within a population, single molecule experiments require data from hundreds of enzyme turnovers. Collecting enough Current Opinion in Chemical Biology 2009, 13:436–442
data for good statistics can be tedious and is facilitated by methods for interrogating large numbers of single molecules at the same time, that is, single molecule multiplexing. Traditionally, multiplexing has been achieved by immobilizing enzymes or substrates (e.g. DNA) on a surface and using conventional and/or fluorescence microscopy to image the molecules (see [1–4] for recent reviews of single molecule optical and force techniques). Recently, a number of new methods for multiplexing have been developed and these are described in Box 1. Several of these methods avoid direct enzyme or substrate immobilization and are likely to increase the number of systems amenable to single molecule techniques. Recent reviews have covered the extensive single molecule literature on systems such as polymerases, ribosomes, helicases and motor proteins [5–8]. This review focuses on recent work on soluble enzymes with small, soluble substrate; highlights important new single molecule work on HIV reverse transcriptase and pays particular attention to the emerging single molecule literature on how peripheral membrane proteins interact with lipid bilayers.
Soluble enzyme cooperativity and heterogeneity b-galactosidase, Inhibition & Enzyme Heterogeneity
A relevant question for oligomeric enzymes is the role of allostery in enzyme activity and inhibition. Escherichia coli b-galactosidase is only active as a tetramer, and the four active sites turnover substrates independently [9,10,11]. One might, therefore, expect that in the presence of inhibitors single b-galactosidase molecules would have 4 distinct activity levels corresponding to 0, 1, 2 or 3 bound inhibitors. However, individual b-galactosidase enzymes exhibited cooperative inhibitor binding and release, switching between states with 0 or 4 bound inhibitors [10]. One surprise was that the activity of single b-galactosidases often significantly changed after inhibitor release, a sign of dynamic heterogeneity caused by protein conformational changes [10]. b-galactosidase dynamic heterogeneity was previously quantified in experiments where individual enzymes immobilized on beads exhibited both long and short waiting times between substrate turnovers [9]. By contrast, experiments in microchambers (Box 1) show that while b-galactosidase molecules display a large distribution of turnover velocities, the velocity of a single b-galactosidase is stable for hundreds of seconds indicating static heterogeneity [12]. Why are different types of heterogeneity observed? Single turnovers were detected in the bead-based experiments, while in the microchambers the results of www.sciencedirect.com
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multiple turnovers were recorded every 15 s. This suggests that on short timescales at ambient temperatures the conformation of a single b-galactosidase enzyme fluctuates around an average conformation with a relatively well-defined long term catalytic rate. This long term rate can change in response to large perturbations such as inhibitor binding. Adenylate kinase enzyme activity and dynamic heterogeneity. All of the evidence for b-galactosidase conformational heterogeneity is indirect because the fluorescent label is on the substrate. Single pair Fo¨rster resonance energy transfer (spFRET) can be used to directly monitor large, functionally important conformational changes and to assess enzyme conformational heterogeneity. In these experiments, a protein is labeled with both a donor and an acceptor fluorophore and changes in the distance between the two probes alters the fluorescence signal (for a guide to spFRET see [13]). Two recent studies of adenylate kinase (AK) used spFRET to measure AK conformational distributions and time-resolved conformational changes in the presence and absence of substrate analogues [14,15]. AK catalyzes the reversible conversion of ATP/AMP to two ADP molecules helping to maintain the energy balance in cells. The AK active site has two flexible lids, the ATP lid in one domain and the AMP lid in the second domain [15]. SpFRET as well as X-ray crystal structures reveal that apo-AK can sample a large distribution of lid conformations, from fully opened to fully closed [14,15]. In the presence of substrate analogue or inhibitor the spFRET distributions shifted to favour the closed conformation [14,15]. For E. coli AK this shift is caused by a doubling of the closing rate constant biasing the conformational distribution towards lid closure in the presence of AMP-PNP, substrate analogue and AMP, substrate [14], thus, demonstrating the utility of spFRET for
directly monitoring dynamic equilibria and the shifting of equilibria by substrates or ligands. AAA+-ATPase Mechanisms Members of the AAA+ ATPase superfamily are generally ring-shaped multisubunit complexes many of which couple ATP binding and hydrolysis to mechanical work. Systems studied using single molecule techniques include F1-ATPase [16,17,18,19,20,21,22], proteins involved in homologous recombination such as helicases [23], RecA [23– 25] and Rad51 [23,26] and the Clp family of unfoldases [27,28]. Recent, elegant single molecule experiments on F1-ATPase, the soluble rotary motor from the F1F0-ATP synthase that can either hydrolyze or synthesize ATP depending on the direction of rotation [29,30], have filled in some of the missing steps in the F1-ATPase mechanism. The minimal F1-ATPase functional complex consists of a g subunit, the rotor, inserted into the centre of the pseudohexameric a3b3 cylindrical stator composed of alternating a and b subunits. Attachment of a filament or beads to the g subunit allows rotation of individual F1ATPase complexes to be observed using conventional or fluorescence microscopy. Such experiments revealed that rotation of the rotor occurs in 1208 steps [31], and these steps have since been shown to consist of 808 and 408 substeps [16,32] (Figure 1). In general, one of the b subunits is empty, bE, one has ATP bound, bATP, and the third has ADP or ADP-Pi bound, bADP [33,34]. ATP binding to bE leads to ADP release from bADP, and the 808 substep during which ATP in the third subunit is hydrolyzed to ADP-Pi [16]. The temperature dependence of F1-ATPase stepping recently revealed that ADP release is rate limiting [17,22]. What about the 408 substep? Excess Pi in solution can drive 408 rotations in the opposite direction showing that Pi release drives the 408 substep [16]. In addition, fluorescently labeled
Box 1 Immobilizing or encapsulating single molecules To ensure that an individual molecule stays in the observation volume, time resolved single molecule microscopy requires limiting the volume available to each molecule. This has often been accomplished using immobilization. However, concerns about the effects of surfaces on enzyme activity as well as the difficulties inherent in selective immobilization have led to the development of encapsulation strategies. Immobilization or encapsulation also allows single molecules to be arrayed so that data from multiple molecules may be collected simultaneously helping to ensure good experimental statistics. A number of immobilization and encapsulation strategies are illustrated. (a) Single molecule experiments often use immobilized proteins (top left) or substrates (top right). (b) Immobilization of large numbers of DNA strands near a nanoscale barrier can be achieved by combining lipid attached DNA molecules (green on orange circles), nanobarriers and flow [26,49]. (c) In this scheme, the lipids diffuse towards the barrier and flow stretches the DNA creating a ‘DNA curtain’. Figure adapted from Fazio et al. [49]. Alternatively, enzymes may be encapsulated in microchambers at the ends of optical fibres, www.sciencedirect.com
where the illustrated reaction is for b-galactosidase hydrolyzing resorfin-b-galactopyranoside. There are hundreds of fibres in the bundles allowing multiplexing, but buffer exchange is complicated. This figure was adapted from Gorris et al. [10]. (d) Enzymes and large substrates such as DNA may be encapsulated in vesicles made porous by incorporating protein pores (left) or bad lipid packing (right). The vesicles are easily immobilized, and the holes in the membrane allow buffer exchange. This figure was adapted from Cisse et al. [24]. (e) Enzymes may be encapsulated in naturally porous virus capsids that can then be immobilized. E is the encapsulated enzyme, S and P, the substrate and product, respectively, can diffuse in and out of the capsid. This figure was adapted from Comellas-Aragones et al. [50]. (f) Enzymes may be encapsulated in aqueous droplets and, as shown in the time series from left to right, the droplets can be fused allowing buffer exchange [51]. The scale bar in the first frame is 1 mm (picture courtesy of LS Goldner). Droplets can also be multiplexed using a number of microfluidic architectures. Current Opinion in Chemical Biology 2009, 13:436–442
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ATP bound at 08 is released as ADP following a 2408 rotation (two 1208 steps). How is the energy gained from the chemical steps converted to mechanical work? Single molecule fluorescence polarization experiments have correlated nucleotide binding states with opening and closing of labeled b subunits suggesting that bending motions of the b subunits drive rotation [20] (Figure 1). It has also been suggested that interactions between the g rotor and both the bottom and top of the a3b3 stator drive rotations [18,19,35]. However, truncated g rotors that only sit on the top of the stator can still generate torque and processively rotate in the correct direction suggesting that biased diffusion may be important for F1-ATPase rotary Current Opinion in Chemical Biology 2009, 13:436–442
motion [18,19]. While more experiments on the details of the mechanical couplings remain to be done, single molecule F1-ATPase experiments are close to providing a complete, detailed molecular mechanism for the conversion of energy from chemical reactions to rotary motion.
Motion on a track and HIV reverse transcriptase (RT) To catalyze reactions on DNA or RNA enzymes must partition from solution unto quasi-one dimensional oligonucleotides and find their targets. Single molecule studies using fluorescently labeled, surface immobilized oligonucleotides and fluorescently labeled proteins have been particularly valuable for elucidating how enzymes bind to, travel on and manipulate oligonucleotides www.sciencedirect.com
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Figure 1
The proposed mechanism for F1-ATPase rotation. ATP binding to the open, empty b subunit, b1, triggers closing of b1. This closing leads to ADP release from the b2 subunit, located 1208 to the left of b1, and an 808 counter-clockwise substep by the g subunit (yellow teardrop). Subsequently inorganic phosphate, Pi, is released from b2 triggering the 408 substep and ATP is hydrolyzed by b3 allowing b3 to partially open. It is currently unclear whether hydrolysis of ATP and the associated b subunit conformational changes occur before or after the rotational substeps. Open, partially closed and fully closed b subunit conformations are shown schematically in red, green and blue, respectively. T indicates bound ATP and D indicates bound ADP. This figure is adapted from Masaike et al. [20].
[5,23,36]. In the past year, spFRET studies by Zhuang and co-workers have provided important insights into how labeled RT interacts with labeled oligonucleotides and how inhibition by noncompetitive non-nucleoside RT inhibitors alters these interactions [37,38]. Heterodimeric RT, which has both DNA polymerase and RNAse H activity, transcribes the viral genome from single stranded RNA to double stranded DNA allowing infected cells to replicate the virus [39]. Intriguingly, the distribution of RT locations on primer-template oligonucleotides as well as RT motion depend on the composition of the oligonucleotide hybrids [37,38]. On mimics for plus strand elongation RT predominantly bound in a polymerase competent manner with the fingers domain at the 30 end of the primer strand [37] (Figure 2). By contrast, RT predominantly bound in the opposite conformation, with the RNase H domain at the 30 end of the primer, for mimics of RNase H substrates. Experiments on hybrid primer strands composed of both RNA and DNA, demonstrate that the 50 composition of the primer strand determines the binding orientation. Thus, the binding orientation and position of RT help determine its function. How does RT find the primer ends and how does it assume the correct orientation? Time-resolved spFRET studies of RT–oligonucleotide interactions show that RT motion is rarely unidirectional and the protein slides back and forth along the track [38]. Sliding is integral to the RT mechanism and elongating RT slides back and forth even when it must displace an RNA strand. Nucelotides and inhibitors alter the dynamic equilibrium for sliding with the rate constant for sliding to the 50 end of the primer (kback) decreasing in the presence of cognate dNTPs and increasing in the presence of the non-nucleoside RT inhibitor nevirapine. In addition to sliding, RT can flip around the substrate to assume the correct orientation, for www.sciencedirect.com
example, flipping from an RNase H competent conformation with the from RNase H domain at the 30 end of the primer to a polymerase competent conformation with the fingers domain at the 30 end of the primer, while still remaining attached to the substrate [38]. Thus, these elegant experiments reveal the dynamic equilibria that are integral to RT function and how biasing sliding favours elongation or inhibition.
Peripheral membrane proteins and interfacial catalysis Substrate turnover by peripheral membrane proteins depends not only on the details of the chemical mechanism but also on how the enzymes partition to and interact with two-dimensional membranes. Single molecule fluorescence microscopy can directly monitor protein motions at the interface and a related techniques fluorescence correlation spectroscopy (FCS) can monitor the diffusion time scale that changes dramatically when small proteins bind much larger vesicles [40,41–43]. Membrane active enzymes can exhibit ‘substrate dilution inhibition’ where a sharp decrease in enzyme activity is observed as the substrate is diluted in the membrane. For extracellular Bacillus thuringiensis phosphatidylinositol specific phospholipase C (PI-PLC), which catalyzes cleavage of the sn-3-phosphodiester bond in phosphatidylinositol (PI), inhibition can occur even when 40% of the membrane is substrate suggesting the involvement of factors other than substrate scarcity [40]. FCS experiments on PI-PLC reveal that the affinity of B. thuringiensis PI-PLC for lipid vesicles initially parallels enzyme activity and Kd drops an order of magnitude for vesicles containing anionic lipids and 10% phosphatidylcholine (PC), an effector lipid that enhances activity [40]. However, unlike the activity, which is stable up to 50% PC and then precipitously drops, Kd continues to decrease as the Current Opinion in Chemical Biology 2009, 13:436–442
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Figure 2
Experimental setup and conformational distributions for RT spFRET experiments. (a) The substrate consisting of an acceptor labeled template strand and a primer strand were immobilized on a quartz coverslip and donor labeled RT was added. The donor and acceptor fluorescence were excited using total internal reflection fluorescence (TIRF) microscopy, which preferentially excites fluorophores near the surface. RT binding to the substrate leads to spFRET and acceptor fluorescence. H indicates the RT RNase H domain and F indicates the fingers domain. The 50 ends of the oligonucleotides are indicated by circles and the 30 ends are indicated by arrows. (b) When binding to plus strand synthesis mimics, DNA(primer)DNA(template), RT primarily binds in a polymerase competent orientation with the fingers domain near the 30 end of the primer resulting in high spFRET efficiencies when the donor is in the RNase H domain (top). RT binds RNA(primer)-DNA(template) substrates in the opposite orientation with the RNase H domain at the 30 end of the primer and low spFRET efficiency (bottom). DNA is shown in black, RNA is shown in red and the donor and acceptor fluorophores are shown in green and red, respectively. Figures are adapted from Abbondanzieri et al. [37].
PC content is increased to 90% indicating tight membrane binding. Parallel NMR studies of the phospholipid headgroups reveal that tight PI-PLC binding preferentially restricts the mobility of the PC headgroup [40]. Thus, substrate dilution inhibition likely arises from substrate scarcity as well as restrictions on protein and lipid mobility. Interactions of fluorescently labeled proteins with supported lipid bilayers and multilayers can be directly imaged with single molecule fluorescence microcopy [44,45,46]. Studies by Yang and co-workers on cobra phospholipase A2 (PLA2), a Ca2+-dependent phospholipase that catalyzes hydrolysis of the sn-2 ester of glycerophospholipids, recently demonstrated that binding and activity preferentially occur near membrane defects [44]. Similar experiments performed by Uji-i and co-workers on a mutant phospholipase A1 (PLA1) that catalyzes hydrolysis of the sn-1 ester of phospholipids also showed preferential binding and activity at defects [45]. Why this preference for defects? High membrane curvature associated with defects or lipid tubes on the top of multilayers probably increase substrate accessibility reducing the energy needed to pull the phospholipid headgroup into the active site [44,45]. PLA1 also showed a number of different interactions with the membranes ranging from fast, transient association to slower diffusion on top of multilayers to occasional immoCurrent Opinion in Chemical Biology 2009, 13:436–442
bilization. The diffusion of single active PLA1 molecules changed over time, transitioning from slow diffusion near membrane edges, probably associated with catalysis, to faster diffusion presumably associated with the search for a new substrate lipid [45]. Inactive PLA1 diffusion was less heterogeneous, generally displaying a diffusion coefficient similar to that expected for a lipid [45]. Similar lipid assisted scooting, where protein motion results from diffusion of a bound lipid, seems to be a general feature of protein–lipid interactions and has been observed in single molecule fluorescence studies of acyl-CoA binding protein bound to POPC giant unilamellar vesicles [47] and for the GRP1 pleckstrin homology domain on supported lipid bilayers [48]. Single molecule experiments have thus begun to correlate different modes of protein–membrane interactions with the catalytic cycle and to visualize binding hotspots associated with enzyme activity.
Conclusions Recent single molecule enzyme studies demonstrate the importance of these techniques for probing catalytically relevant conformational changes, the cooperativity of activity and inhibitor binding/release in oligomeric proteins as well as probing interaction between proteins and oligonucleotides or membranes. As highlighted by the work on b-galactosidase, RT and PI-PLC, single fluorescence molecule techniques are particularly useful for determining the molecular mechanisms of inhibition www.sciencedirect.com
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in complicated systems. The advent of new methods for protein encapsulation as well as microfluidic methods may obviate the need for surface immobilization, expanding the applicability of these methods.
Acknowledgements Research in the author’s laboratory is funded in part by a grant from the Alpha-1 Foundation and supported by a charitable donation from Talecris Biotherapeutics, Center for Science and Education.
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