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What type of animation software is used to create these molecular motor motions?

What type of animation software is used to create these molecular motor motions?


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Recently, I came across this video on molecular motor motion. Can someone suggest software that can do similar animations? I am not sure what they (at Harvard Medical School) used but the animations are really cool.


Most any reasonable animation/modeling software can handle this sort of animation. I have developed an educational iOS app with interactive biological macromolecules, including several animations. Here is how I would approach this, I suspect something similar was done here.

The models are likely built up from PDB files, or equivalent. These files allow viewing and manipulating large molecules in 3D with special viewers. Many 3D modeling/animation apps can also view and convert these molecules into polygonal models that can be used as components and then animated within the app. I use Cinema 4D, not cheap. There is a free, downloadable program called Blender. I'm sure Blender can handle the animation but not certain whether it can read and convert the PDB file. If not, there may be other options for this.

This is not a challenge to be taken lightly. The person doing the animation you reference also knew how to assemble the parts as well as how the movement is effected. There is also a steep learning curve to using this type of software.


RING NMR dynamics: software for analysis of multiple NMR relaxation experiments

Molecular motions are fundamental to the existence of life, and NMR spectroscopy remains one of the most useful and powerful methods to measure their rates and molecular characteristics. Multiple experimental methods are available for measuring the NMR relaxation properties and these can require different methods for extracting model parameters. We present here a new software application, RING NMR Dynamics, that is designed to support analysis of multiple relaxation types. The initial release of RING NMR Dynamics supports the analysis of exponential decay experiments such as T1 and T2, as well as CEST and R2 and R relaxation dispersion. The software runs on multiple operating systems in both a command line mode and a user-friendly GUI that allows visualizing and simulating relaxation data. Interaction with another program, NMRFx Analyst, allows drilling down from the derived relaxation parameters to the raw spectral data.


Assembling Life’s Molecular Motor

Despite the grand diversity among living organisms, the molecule used to store and transmit energy within aerobic, or oxygen-using, cells is remarkably the same. From bacteria to fungi, plants, and animals, adenosine triphosphate (ATP) serves as the universal energy currency of life, fueling the processes cells need to survive and function.

Over the course of a day, an individual will typically use the equivalent of his or her bodyweight in ATP however, the human body carries only a small amount of the molecule at any one time. That means cells must constantly recycle or replenish their limited capacity, relying on a highly efficient molecular motor called ATP synthase to do the job.

As part of a project dedicated to modeling how single-celled purple bacteria turn light into food, a team of computational scientists from the University of Illinois at Urbana-Champaign (UIUC) simulated a complete ATP synthase in all-atom detail. The work builds on the project’s first phase—a 100-million atom photosynthetic organelle called a chromatophore—and gives scientists an unprecedented glimpse into a biological machine whose energy efficiency far surpasses that of any artificial system.

First proposed under the leadership of the late Klaus Schulten, a pioneer in the field of computational biophysics and the founder of the Theoretical and Computational Biophysics Group at UIUC, the research has progressed under the stewardship of Abhishek Singharoy, co-principal investigator and a National Science Foundation postdoctoral fellow with UIUC’s Center for the Physics of Living Cells.

In addition to Singharoy, the team includes members from the groups of UIUC professors Emad Tajkhorshid, Zaida Luthey-Schulten and Aleksei Aksimentiev research scientist Melih Sener and developers Barry Isralewitz, Jim Phillips, and John Stone. Experimental collaborator Neil Hunter of the University of Sheffield in England also took part in the project.

The UIUC-led team built and tested its mega-model under a multiyear allocation awarded through the Innovative and Novel Computational Impact on Theory and Experiment program on the Titan supercomputer, a Cray XK7 managed by the US Department of Energy’s (DOE’s) Oak Ridge Leadership Computing Facility (OLCF), a DOE Office of Science User Facility located at DOE’s Oak Ridge National Laboratory.

Using Titan, the team produced a virtual tool that can predict in exacting detail the chemical energy output of a photosynthetic system based on the amount of sunlight absorbed. The research could one day contribute to advanced clean energy technology that incorporates biological concepts.

“Nature has designed the chromatophore in such a way that it can generate enough ATPs for these bacteria to survive in low-light environments such as the bottom of ponds and lakes,” Singharoy said. “Our work captured this energy conversion process in all-atom detail and allowed us to predict its efficiency.”

Light in motion

Often referred to as the power plant of the cell, ATP synthase is a complex enzyme that speeds up the synthesis of its molecular precursors, adenosine diphosphate (ADP) and phosphate. Embedded within the chromatophore’s inner and outer membrane, the enzymatic motor consists of three major parts—an ion-powered rotor, a central stalk, and a protein ring.

Similar to a waterwheel that’s turned by the force of a flowing stream, the ATP synthase rotor harnesses the electrochemically spurred movement of ions, such as protons or sodium, from high concentration to low concentration across the membrane. The resulting mechanical energy transfers to the central stalk, which assists the protein ring in synthesizing ATP.

Remarkably, the process works just as well in reverse. When too many ions build up on the outer side of the chromatophore, the ATP synthase protein ring will break down ATP into ADP, a process called hydrolysis, and ions will flow back to the inner side.

“Normally, you would expect a lot of energy loss during this process, like in any man-made motor, but it turns out ATP synthase has very little waste,” Singharoy said. “How this motor is designed to minimize energy loss is the question we started asking.”

Similar to a tinkerer disassembling an engine to better understand how it works, Singharoy’s team broke down the 300,000-atom enzyme into its constituent parts. Drawing from decades of research into ATP synthase, past models, and new experimental data supplied by a Japanese team led by Takeshi Murata of the RIKEN Center for Life Science Technologies, the team constructed and simulated the pieces of the ATP synthase puzzle independently and together on Titan.

To capture important processes that play out over millisecond time scales, Singharoy, in collaboration with Christophe Chipot of the French National Center for Scientific Research and Mahmoud Moradi of the University of Arkansas, deployed the molecular dynamics code NAMD strategically. The team executed an ensemble strategy, tracking the motion of around 1,000 replicas of ATP synthase simultaneously with time steps of 2 femtoseconds, or 2,000 trillionths of a second. In total, the team accumulated 65 microseconds (65 millionths of a second) of simulation time, using this information to extrapolate motions that occur over the course of a millisecond (1 thousandth of a second).

As a result, the team identified previously undocumented swiveling motions in the protein ring that help explain the molecular motor’s efficiency. Similarly, the team’s simulations captured the rubber band–like elasticity of the enzyme’s central stalk. Singharoy’s team estimated that when paired with the protein ring, the stalk absorbs about 75 percent of the energy released during hydrolysis.

Additionally, simulations of the protein ring by itself revealed a unit that can function independently, a finding reported in experiments but not in computational detail. “Even in the absence of the center stalk, the protein ring itself is capable of ATP hydrolysis. It’s not very efficient, but it has the capability,” Singharoy said.

The big picture

After simulating its complete ATP synthase model, the UIUC team incorporated the enzyme into its previously constructed chromatophore model to gain the most comprehensive picture of a photosynthetic system to date.

With this virtual biological solar panel, the team could measure each step of the energy conversion process—from light harvesting, to electron and proton transfer, to ATP synthesis—and better understand its mechanical underpinnings.

Nature’s chromatophore is designed for low-light intensity, only absorbing between 3 and 5 percent of sunlight on a typical day. The team, through the efforts of Sener, found this absorption rate translates to around 300 ATPs per second, which is the amount a bacterium needs to stay alive.

Having studied nature’s design, the team now wanted to see if it could improve upon it. Assuming the same amount of light intensity, the team designed an artificial chromatophore with a decidedly unnatural protein composition, boosting the presence of two types of specialized proteins. Analysis of the new design predicted a tripling of the photosynthetic system’s ATP production, opening up the possibility for the chromatophore’s human-guided optimization.

“You could potentially genetically modify a chromatophore or change its concentration of proteins,” Singharoy said. “These predictions promise to bring forth new developments in artificial photosynthesis.”

Under its latest INCITE allocation, the UIUC team is pivoting to energy conversion in a different lifeform: animals. Taking what it has learned from modeling photosynthesis in purple bacteria, the team is modeling cellular respiration, the process animal cells use to convert nutrients to ATP.

“You have at least two proteins in common between respiration and photosynthesis,” said Singharoy, who is continuing his involvement with the project as an assistant professor at Arizona State University. “The question is what design principles carry over into higher organisms?”

Life in situ

Simulation of the chromatophore—complete with ATP synthase—marks an ongoing shift in computational biophysics from analyzing individual cell parts (e.g., single proteins and hundreds of atoms) to analyzing entire cell systems (e.g., hundreds of proteins and millions of atoms).

Schulten, who passed away in October 2016, understood better than most people the significance of using computers to simulate nature. In an interview in 2015, he laid out his rationale for modeling the chromatophore. “The motivation is to understand a very key step of life on Earth on which all life depends today. Energy-wise 95 percent of life on Earth depends on photosynthesis, including humans,” he said.

Schulten also understood the milestone a specialized organelle represented on the road to simulating a complete single-celled organism. “We don’t have anything smaller than a cell that we would call alive,” he said. “It’s the smallest living entity, and we want to understand it.”

With next-generation supercomputers, including the OLCF’s Summit, set to come online in 2018, the research group Schulten founded in 1989 is preparing to take on the grand challenge of simulating a cell.

Under the leadership of Tajkhorshid, the team plans to simulate the first billion-atom cell, including the basic components a cell needs to survive and grow. Improvements to NAMD and work being done under the OLCF’s Center for Accelerated Application Readiness program are helping to make the vision of Schulten and others a reality.

“We keep moving forward,” Singharoy said. “Our exhaustive study of a complete organelle in all-atom detail has opened the door for a full cell in all-atom detail.”

Abhishek Singharoy, Christophe Chipot, Mahmoud Moradi, and Klaus Schulten, “Chemomechanical Coupling in Hexameric Protein–Protein Interfaces Harness Energy Within V–Type ATPases.” Journal of the American Chemical Society 139, no. 1 (2017): 293–310, doi:10.1021/jacs.6b10744.

Abhishek Singharoy, Angela M. Barragan, Sundarapandian Thangapandian, Emad Tajkhorshid, and Klaus Schulten, “Binding Site Recognition and Docking Dynamics of a Single Electron Transport Protein: Cytochrome c2.” Journal of the American Chemical Society 138, no. 37 (2016): 12077–12089, doi:10.1021/jacs.6b01193.

Melih Sener, Johan Strumpfer, Abhishek Singharoy, C. Neil Hunter, and Klaus Schulten, “Overall Energy Conversion Efficiency of a Photosynthetic Vesicle.” eLife 5 (2016): e09541, doi:10.7554/eLife.09541.


Contents

The National Highway Traffic Safety Administration funded the first national guidelines for the standardization training in the field of traffic collision reconstruction in 1985. This led to the establishment of "Accreditation Commission for Traffic Accident Reconstruction" (ACTAR), an industry accreditation group. [1] This field of motorcycle collision research was pioneered by Hugh H. Hurt Jr. His meticulous collision reconstructions of motorcycle collisions helped to explain that proper helmets reduced head injuries, most motorcyclists needed more driver training to control skids, and a large percentage of motorcycle collisions involved left-turning automobiles turning in front of the oncoming motorcycle. [2]

Scene inspections and data recovery involves visiting the scene of the collision and investigating all of the vehicles involved in the collision. Investigations involve collecting evidence such as scene photographs, video of the collision, measurements of the scene, eyewitness testimony, and legal depositions. Additional factors include steering angles, braking, use of lights, turn signals, speed, acceleration, engine rpm, cruise control, and anti-lock brakes. Witnesses are interviewed during collision reconstruction, and physical evidence such as tire marks are examined. The length of a skid mark can often allow calculation of the original speed of a vehicle for example. Vehicle speeds are frequently underestimated by a driver, so an independent estimate of speed is often essential in collisions. Inspection of the road surface is also vital, especially when traction has been lost due to black ice, diesel fuel contamination, or obstacles such as road debris. Data from an event data recorder also provides valuable information such as the speed of the vehicle a few seconds before the collision. [3]

As part of the investigation of a vehicle collision, an investigator typically documents evidence at the collision site and the damage to the vehicles. The use of 3-dimensional laser scanning has become a common method for documentation. The product of scanning is a 3D point cloud that can be used to take measurements and create computer models used in the analysis of the collision. The 3D data can be incorporated into many of the computer simulation programs used in collision reconstruction. The 3D point clouds and models can also be used for creating visuals to illustrate the analysis and to show views of witnesses and the involved drivers.

Many new vehicles are equipped with onboard "Crash Data Recorders or Event Data Recorders" (CDR or EDR). The Bosch CDR Tool is a commercially available tool, allowing to image crash data directly from all supported vehicles giving a detailed report of critical data parameters leading up to and during a crash. Some of the parameters include pre-crash data, vehicle speed, brake status, throttle position, ignition cycles, delta-V, seat belt status, and others. [4]

Hyundai and Kia as well as most heavy commercial vehicles are equipped with EDR, however are not supported by Bosch equipment. To access this information a diagnostic retrieval tool unique to these manufacturers is required.

Vehicular collision reconstruction analysis includes processing data collecting, evaluating possible hypotheses, creating models, recreating collisions, testing, and utilizing software simulations. Like many other technical activities, collision reconstruction has been revolutionized by the use of powerful, inexpensive computers and specialty software. Various types of collision reconstruction software are used to recreate crash and crime scenes and to perform other useful tasks involved in reconstructing collisions. Collision reconstruction software is regularly used by law enforcement personnel and consultants to analyze a collision and to demonstrate what occurred in a collision. Examples of types of software used by collision reconstructionists are CAD (computer aided design) programs, vehicle specification databases, momentum and energy analysis programs, collision simulators, and photogrammetry software.

After the analysis is completed, forensic engineers compile report findings, diagrams, and animations to form their expert testimony and conclusions relating to the collision. Forensic animation typically depicts all or part of a collision sequence in a video format so that non-technical parties, such as juries, can easily understand the expert's opinions regarding that event. To be physically realistic, an animation needs to be created by someone with a knowledge of physics, dynamics and engineering. When animations are used in a courtroom setting, they should be carefully scrutinized. Animation software can be easily misused, because motions which are not physically possible can be displayed. A reliable animation must be based on physical evidence and calculations which embody the laws of physics, and the animation should only be used to demonstrate in a visual fashion the underlying calculations made by the expert analyzing the case. [5]

Motorcycle collision reconstruction is similar to other collision reconstruction techniques and relies on the same basic principles of conservation of energy and momentum as automobile collision reconstruction plus adds the specifics of motorcycle dynamics and rider control. Proper reconstruction of a motorcycle collision requires detailed knowledge of motorcycle dynamics plus knowledge of how motorcycles react to rider input.

Motorcycle collision reconstruction follows reverse a chronological order of events, working from the point of rest of the motorcycle and/or rider backwards to a point in time before to the start of the collision sequence to when possible actions could have prevented the crash.

Motorcycle collision reconstruction relies on knowledge of the five phases of a motorcycle collision.

Perception–reaction: This is the phase where the rider perceives a collision hazard and decides on a response. Perception/reaction time is estimated at 1.1 to 1.5 seconds. [6]

Avoidance – braking/steering: In this next phase, the rider typically engages in some type of avoidance using steering or braking using the front brake, rear brake or a combination. Physical evidence at the scene combined with statements from witnesses can give clues as to what type of avoidance occurred.

Pre-impact sliding: During braking, riders may overuse the motorcycle brakes, resulting in locking the front and/or rear wheel. If the front wheel locks, the rider will almost certainly lose control and crash. If the rider loses control and crashes while braking, the motorcycle and rider usually separate and slide in the same trajectory they were moving in before the crash.

Impact: The bike and/or rider may collide with other object like a vehicle or guardrail. Damage caused by impact can be evaluated and combined with sliding distance to help determine the motorcycle's speed during the collision sequence.

Post-impact motion: After impact, additional movement to the point of final rest can occur. The rider frequently separates from the motorcycle and travels independently to the final point of rest. Analysis of post-impact travel distance can also determine speeds associated with the collision. [7]

The Royal Canadian Mounted Police conducts On-Scene Collision Investigation (Level-2), Advanced Collision Analysis (Level-3), and Forensic Collision Reconstruction (Level-4) as well as Commercial Vehicle Collision and Pedestrian/Bicycle Collision courses at the Pacific Region Training Center (PRTC) located in Chilliwack, British Columbia. These courses are also available to Non-RCMP Police Agencies. [8]

Northwestern University Center for Public Safety conducts Traffic Crash Investigation courses utilized by both law enforcement and public agencies. [9]

The Institute of Police Technology and Management (IPTM) is a recognized institute for Crash Investigation for Law Enforcement as well as professional agencies. [10]

The Royal Canadian Mounted Police utilize full-time Forensic Collision Reconstructionists and Analysts as a service line. In British Columbia, they are referred to as ICARS (Integrated Collision Analysis and Reconstruction Service). [11] ICARS units are located in each RCMP District within the Province of B.C.

California Highway Patrol utilize a team deployment called MAIT ("Multidisciplinary Accident Investigation Team"). Each team consists of inspectors with specialized training in traffic collision reconstruction, traffic engineering, automotive engineering, and vehicle dynamics. MAITs are composed of one CHP sergeant (the team leader), two or more CHP officers, one Motor Carrier Specialist I (MCS I), and one Senior Transportation Engineer from Caltrans. [12]


Biologically inspired artificial muscles made from motor proteins

Inside our cells, and those of the most well-known lifeforms, exist a variety of complex compounds known as "molecular motors." These biological machines are essential for various types of movement in living systems, from the microscopic rearrangement or transport of proteins within a single cell to the macroscopic contraction of muscle tissues. At the crossroads between robotics and nanotechnology, a goal that is highly sought after is finding ways to leverage the action of these tiny molecular motors to perform more sizeable tasks in a controllable manner. However, achieving this goal will certainly be challenging.

"So far, even though researchers have found ways to scale up the collective action of molecular motor networks to show macroscopic contraction, it is still difficult to integrate such networks efficiently into actual machines and generate forces large enough to actuate macroscale components," explains Associate Professor Yuichi Hiratsuka from the Japan Advanced Institute of Science and Technology, Japan.

Fortunately, Dr. Hiratsuka, in collaboration with Associate Professor Takahiro Nitta from Gifu University and Professor Keisuke Morishima from Osaka University, both in Japan, have recently made remarkable progress in the quest to bridge the micro with the macro. In their latest study published in Nature Materials, this research team reported the design of a novel type of actuator driven by two genetically modified biomolecular motors. One of the most attractive aspects of their biologically inspired approach is that the actuator self-assembles from the basic proteins by simple light irradiation. In a matter of seconds after light hits a given area, the surrounding motor proteins fuse with rail-like proteins called microtubules and arrange themselves into a hierarchical macroscopic structure that resembles muscle fibers.

Upon formation around the target (illuminated) zone, this "artificial muscle" immediately contracts, and the collective force of the individual motor proteins is amplified from a molecular scale to a millimeter one. As the scientists showed experimentally, their approach could be ideal for small-scale robotics applications, such as actuating microscopic grippers to handle biological samples (Figure 1). Other millimeter-scale applications also demonstrated include joining separate components together, such as miniature cogwheels, and powering minimalistic robotic arms to make an insect-like crawling microrobot.

What's also very remarkable about this technique is that it is compatible with existing 3D printing techniques that use light, such as stereolithography. In other words, microrobots with built-in artificial muscles may be 3D printable, enabling their mass production and hence increasing their applicability to solve various problems. "In the future, our printable actuator could become the much-needed 'actuator ink' for the seamless 3D printing of entire robots. We believe that such a biomolecule-based ink can push forward the frontier of robotics by enabling the printing of complex bone and muscle components required for robots to further resemble living creatures," says Dr. Hiratsuka.

One potential improvement to the present technique would be finding ways to efficiently decontract the artificial muscles (reversibility). Alternatively, the present strategy could also be changed so as to produce spontaneous oscillatory behavior instead of contraction, as is observed in the mobile cilia of microbes or in insect flight muscles.

In any case, this study effectively shows how mimicking the strategies that nature has come up with is often times a recipe for success, as many scientists in the field of robotics have already figured out.


Motor Molecules Use Random Walks To Make Deliveries In Living Cells

Cells rely on tiny molecular motors to deliver cargo, such as mRNA and organelles, within the cell. The critical nature of this transport system is evidenced by the fact that disruption of motors by genetic defects leads to fatal diseases in humans. Although investigators have isolated these motor to study their function in a controlled environment outside the cell, it has been difficult for researchers to follow these fascinating molecular transporters in their natural environment, the living cell.

Now, two articles published by Cell Press in Biophysical Journal, make use of incredibly tiny, glowing "quantum dots" to track the miniscule motions of myosin V in living cells. Interestingly, both research groups independently report that myosin V molecules carry their quantum dot cargo either in a straight line or in a manner akin to a drunken walk.

Myosin V is a motor molecule that "walks" in a fashion similar to humans by stepping along actin filament tracks that are assembled in a dense, criss-crossing network inside the cell. A critical feature of these motors is their ability to walk long distances without falling off their tracks. However, this has never been observed within cells. Through the binding of quantum dots directly to a single myosin V molecule, both investigative teams used sophisticated microscopes and sensitive cameras to witness the 72 nanometer strides (equivalent to 1 millionth of an inch) taken by these motors for the first time in cells.

In results published in the May 20th 2009 issue of Biophysical Journal, Dr. Giovanni Cappello from the Institut Curie in Paris, France tracked the movement of single myosin V molecules with inside living HeLa cells. Dr. Cappello and colleagues reported that the myosin V can transport cargo for long distances without falling off its track at velocities higher than would be expected based on earlier studies. "Our approach goes beyond conventional experiments on organelles and opens interesting perspectives for studying intracellular transport pathways and how motors behave in complex filament networks," says Dr. Cappello.

Dr. David Warshaw and colleagues from the University of Vermont College of Medicine used quantum dots to follow the activity of myosin V in COS-7 cells. Their findings, published in the July 22nd 2009 issue of the journal, suggested that myosin V's apparent drunken walk is in fact the motor taking turns at almost every intersection it encounters along the dense and randomly oriented intracellular actin highway. "Cargo delivery in cells can't totally be a random process, therefore, using the approach described here we can characterize how motors and cargo link up and understand the engineering design principles Mother Nature uses to guarantee efficient and effective delivery of cargo within cells," offers Dr. Warshaw.

Researchers include Shane R. Nelson, M. Yusuf Ali, Kathleen M. Trybus, David M. Warshaw, of the department of Molecular Physiology and Biophysics, University of Vermont College of Medicine, Burlington, VT.

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Materials provided by Cell Press. Note: Content may be edited for style and length.


RESULTS

While cryo-EM has emerged as a powerful technique to obtain near atomic resolution structures of large complexes (37, 38), it is still challenging to resolve inherently flexible components (e.g., molecular motors) to high resolution. Further, properly reconstructing any symmetry mismatches between the massive capsid and relatively small motor is still nontrivial (4, 26, 29, 3941). Hence, the most tractable path to the atomic structure of a functional viral dsDNA packaging motor is to determine atomic resolution structures of isolated components and then fit them into cryo-EM reconstructions of the entire motor complex assembled on the procapsid (42). To date, we have determined atomic resolution structures of nearly every component of the phi29 packaging motor including the portal (25), the prohead-binding domain of the pRNA (27), the pRNA CCA bulge (43), part of the A-helix of the pRNA (27, 44, 45), the N-terminal ATPase domain of gp16 (31), and, most recently, the C-terminal vestigial nuclease domain of gp16 (46). Further, we have been recently determining the full-length structure of a homologous packaging ATPase from a relative of phi29, bacteriophage asccphi28 (47).

Cryo-EM reconstruction of particles stalled during DNA packaging

Having completed the library of atomic resolution structures of phi29 motor components, we turned to single-particle cryo-EM to image functional motors assembled on procapsids (fig. S1). Briefly, we incubated 120-base pRNA proheads with gp16, phi29 genomic DNA, ATP, and magnesium for approximately 5 min before stopping the packaging reaction via addition of an excess of the nonhydrolysable ATP analog ATP-γ-S. This incubation period is long enough to allow gp16 to assemble on proheads and for functional motors to fully package their genomic DNA substrates. The addition of ATP-γ-S serves two purposes: (i) It stabilizes packaged particles such that the motor remains attached, and the DNA is retained in heads and (ii) it drives the motor toward a uniform, substrate-bound state. Presumably, addition of ATP-γ-S arrests the motors near the end of the dwell phase, when all five subunits are bound to substrate (or nonhydrolysable substrate analog ATP-γ-S), and the motor is poised to enter the burst (Fig. 1B, double asterisk).

Procapsid structure

Particles were initially reconstructed assuming C5 symmetry. The resulting map was of high quality (Fig. 2, A to C), as evidenced by readily recognizable secondary structures and often clear and unambiguous side-chain density (Fig. 2C). The overall resolution of the map is

2.9 Å as estimated by “gold standard” Fourier shell correlation (FSC) between independently processed half-datasets (48, 49). Consistent with previous reconstructions of phi29 particles (50, 51), the capsid appears to have pseudo-D5 symmetry that is broken by the unique motor vertex at one end of the otherwise T = 3, Q = 5 prolate icosahedral shell. The density for the capsid was particularly good, nearly the entire length of the capsid protein density could be traced, and side chains were unambiguously positioned (Fig. 2, C and D). Further details of model building and refinement of the procapsid will be described elsewhere, but Fig. 2 (A to D) shows the overall quality of the density corresponding to the protein shell.

(A) Side view of stalled-particle reconstruction, colored by cylindrical radius. Components of the packaging motor are colored magenta. (B) Zoom in on one capsid hexamer, outlined in red in (A). (C) The top panel shows the atomic model of the capsid protein built into density corresponding to one monomer in the hexamer shown in (B). The bottom panel shows the ribbon diagram with the N-terminal HK97 and the C-terminal immunoglobulin (IG)–like domains colored blue and pink, respectively. (D) Cross-section of the reconstruction of stalled particles colored as in (A) but with packaged DNA colored red. (E) Focused C12 reconstruction of the portal. The top panel shows an end-on view with the fitted portal structure (yellow ribbon PDB ID: 1FOU). The lower panel shows a side view of the central helical domain. (F) Focused C1 reconstruction of the entire portal vertex with the atomic model of the portal built into its corresponding density. The top and middle panels show end-on views looking from inside the capsid (upper) and from below the portal (middle) the portal, pRNA, and capsid are shown in yellow, magenta, and blue, respectively. The bottom panel shows the side view. (G) The top panel shows the structure of a fragment of the pRNA (PDB ID: 4KZ2) fitted into its corresponding density. The bottom panel shows the atomic resolution structure of the pRNA built into its corresponding density. The E loop of the pRNA attaches to E loops in the capsid protein, colored blue, green, and red.

Symmetry breaking: Portal structure

As would be expected, the density for the connector was initially poor. Because of the symmetry mismatch between the dodecameric connector and the fivefold symmetric vertex where it sits, imposing C5 symmetry causes the portal to be incoherently averaged around its 12-fold symmetry axis (29). Hence, to improve the density for the portal protein, we used a variation of focused reconstruction and symmetry breaking (Supplementary Materials) (29, 39, 41, 52). This allowed us to obtain a near atomic resolution reconstructions of both the portal alone (Fig. 2E) and the larger asymmetric portal vertex wherein both the 12-fold symmetric portal and the 5-fold symmetric circumscribing capsid proteins are resolved (Fig. 2F).

PRNA structure

Density corresponding to the pRNA in the initial C5 map was not as clear as density corresponding to the capsid protein but was still easily recognizable as RNA (Fig. 2, F and G). Major and minor grooves were apparent with dimensions such as those predicted for A-form nucleic acid helices, and serrated periodicity consistent with the phosphate backbone was visible for much of the pRNA. Density for the 74 bases corresponding to the prohead-binding domain of the pRNA was better than density corresponding to the A-helix, likely due to increased flexibility of the latter. Density corresponding to the pRNA in the C1 map obtained via symmetry breaking based on portal density described above was similar to density in the original C5 map.

Symmetry breaking: pRNA-ATPase structure

Since symmetry breaking based on focused reconstruction of the portal resulted in a high-quality reconstruction of the portal itself (Fig. 2, E and F), it was unexpected that the density corresponding to the ATPase and DNA in this reconstruction was still rather poor and did not seem much improved relative to the original C5 map (Fig. 2D) or to a previously published 12-Å map (31). A possible explanation is that the motor is highly dynamic, and thus, different individual motors are in different conformations and orientations relative to the portal, resulting in smeared density upon interparticle averaging during three-dimensional (3D) reconstruction. Another possibility is that there is yet another symmetry mismatch between the procapsid-portal complex and the ATPase/DNA even once the portal and procapsid are simultaneously aligned, there are still five different ways that the ATPase and DNA can be arranged with respect to the prohead portal.

To test this possibility, we repeated the symmetry-breaking procedure described above, except that, instead of focusing exclusively on the portal, we focused on density that includes the portal, the pRNA, the DNA, and the ATPase. Further, since DNA in the capsid is not well resolved, it is impossible to correctly subtract DNA density from experimental images. Hence, we excluded views where the capsid is projected onto the motor and only included particles where the ATPase was clearly visible. In the resulting reconstruction, the density for the pRNA, DNA, and ATPase were greatly improved in the resulting map (Fig. 3 EMD-22441). While the resulting resolution was good,

4.4 Å overall as estimated by FSC (48, 49) and the D99 criterion (53), density quality varied over the reconstructed volume, and the NTD and associated DNA and pRNA were probably closer to

6-Å resolution and thus not high enough to identify side chains/bases or trace the polypeptide/nucleic acid chain ab initio. Nonetheless, it was sufficient to clearly see secondary structural elements in the ATPase and the major and minor grooves in the DNA and pRNA. The phosphate backbone was still readily apparent in the pRNA density and was now visible in most of the DNA density as well. Density for the portal was not nearly as well resolved as in the previous focused reconstructions of portal-only and the portal-capsid vertex (Fig. 2, E and F, and fig. S2). While density in the radial direction was well resolved, density in circumferential directions was rather poor, indicating that particles were incoherently averaged around the 12-fold symmetry axis of the portal. This result suggests that symmetry breaking was dominated by signal from the pRNA, ATPase, and DNA and that the portal density was not aligned in the selected orientations. This could occur if there is no defined relationship between the portal and the motor. Alternatively, there could be five different ways to arrange the portal with respect to the ATPase, but the small mass of the portal could not provide enough signal to successfully classify these different arrangements. We suspect the latter to be true.

(A) Density for the CTD planar ring viewed looking from below (left) and above (right). Five copies of the CTD NMR structure (PDB ID: 6V1W) are fitted into their corresponding densities and shown as red, yellow, green, orange, and blue ribbon diagrams. (B) Density for the NTD helical assembly viewed looking from below (left) and above (right). Five copies of the NTD crystal structure (PDB ID: 5HD9) are fitted into their corresponding densities and shown as red, yellow, green, orange, and blue ribbon diagrams. (C) A cutaway side view of the phi29 ATPase motor, with CTD and NTD ribbon diagrams colored as above. The pRNA (periphery) and DNA (center) are shown as tan ribbon diagrams in all three panels.

Building an atomic model of the motor

While the separate structures for the prohead binding and portions of the A-helix domains (27, 44, 45) could be fit into their corresponding densities (Fig. 2G), part of these structures had engineered mutations to facilitate crystallization. Hence, since density for the pRNA was of reasonably high quality, a complete 120-base pRNA model was built directly into its corresponding density. However, it proved useful to use base-pairing patterns observed in the crystallographic structures as restraints when refining the model against density. Similarly, an atomic model of the DNA was built by first fitting coordinates of a stretch of ideal B-form DNA into its density. Further, the improved density allowed us to unambiguously fit our crystal structure of the N-terminal ATPase domain [NTD Protein Data Bank (PDB) ID: 5HD9] and our nuclear magnetic resonance (NMR) structure of the C-terminal vestigial nuclease domain (CTD PDB ID: 6V1W) into their corresponding densities (Fig. 3). The linker region between the domains was built directly into density based on the equivalent linker region in our recently determined full-length ATPase structure of a homolog of gp16, the dsDNA packaging ATPase from bacteriophage asccphi28 (

45% similarity to gp16) (47). Fitted structures were refined against their corresponding densities using the real-space cryo-EM refinement module in PHENIX (53). After initial refinement, the map was improved via local averaging over equivalent subvolumes corresponding to the N- and C-terminal domains of the ATPase and to the prohead-binding and A-helix domains of the pRNA. The refined models were then manually adjusted to fit this “locally averaged” map and then refined together against the original, unaveraged map, as in the application of noncrystallographic symmetry averaging in x-ray crystallography.

Gross features of the packaging motor

Several gross features of the motor structure are immediately apparent upon an examination of the refined coordinates (Fig. 4 PDB ID: 7JQQ). First, the pRNA and ATPase are pentameric assemblies, definitively resolving a long-standing controversy regarding the symmetry of these components and refuting several reports of hexameric assemblies (Fig. 4C) (7). Second, as previously reported (54), the pRNA does not substantially interact with the portal but rather bases in the pRNA E loop (U54, G55, and A56) interact with residues in surrounding capsid protein E loops (Gln 117 , Lys 118 , Asp 121 , and Arg 122 of one E loop and residue Gln 117 of a neighboring E loop). Also, consistent with previous results (31), the CTD sits above the NTD and just below the portal where it is wedged between the pRNA and the incoming DNA (Fig. 4B). This arrangement of ATPase domains differs from an arrangement reported for the T4 DNA packaging motor, where structural data suggested a reversed domain polarity wherein the NTD attaches to the portal with the CTD hanging just below (55).

Structure of the packaging motor rendered as molecular surfaces (top panels) and ribbon diagrams (bottom panels) shown from the following: (A) side view (B) cutaway side view to visualize the DNA in the central channel and (C) an end-on view, looking from below the motor. The portal and pRNA are colored cyan and magenta, respectively, and the five ATPase subunits are labeled S1 to S5 and colored blue (S1), orange (S2), green (S3), yellow (S4), and pink (S5). Approximate levels of the CTD and NTD are indicated by blue arrows in the panel. Deviations from the rotational component of helical symmetry are shown indicating loose and tight interfaces by red and green asterisks, respectively, in (C) the loose interfaces are on either side of the lowest subunit in the ATPase helix and correspond to the two active sites where there is no clear density for ATP. Two black lines in the bottom half of (B) are drawn approximately coincident with the helical axis of the DNA before and after the kink that occurs between the CTD and NTD the

12.5° angle between the lines is indicated.

Both the N- and C-terminal domains appear to interact with incoming DNA (Figs. 3 and 4), again differing from results for T4 that suggest that only the CTD interacts with DNA (55). In addition, all five NTDs and CTDs appear to interact with DNA, in contrast to previous results we reported that only one subunit contacts the DNA (35). Further, the DNA structure differs considerably from canonical B-form DNA. It is stretched/partially unwound in some places, compressed in others, and has a prominent kink between the CTD and NTD, deflecting the direction of the DNA

15° from the expected direction of DNA translocation (Figs. 3 and 4, B and C). Hence, mechanisms requiring strict helical symmetry of B-form DNA may need to be reevaluated.

While the pRNA and CTD have approximate C5 symmetry, the NTD of the ATPase is not arranged as a planar ring but rather adopts an arrangement more akin to a cracked ring or a short, one-turn helical assembly (Figs. 4 and 5 and movie S1). The five NTDs in the “helix” roughly wrap one turn (

10 bp) of dsDNA and are thus arranged such that the two helices are approximately in register about every

2 bp. Further, the NTD does not adopt a perfect helical arrangement, as there is some variation in both the twist and rise-relating adjacent subunits (Figs. 4 and 5), with subunits S1 and S5 at the top and bottom of the helix differing most in regard to both twist and rise. Since the NTDs are identical, it is not entirely clear how these deviations arise. A possible partial explanation is that the helical arrangement of NTDs results from subunit interactions with DNA, and thus, the observed distortions in pitch, twist, and direction of the DNA give rise to complementary deviations in the NTD helix. This would explain why S5 deviates most it sits at the bottom of the helix where the DNA is furthest offset. However, it does not explain why S1 deviates, since it is positioned at the top of the helix where the DNA offset is minimal. However, it neighbors S5, and so, the deviation of S5 could influence the position of S1. In addition, while all subunit NTDs contact the pRNA via a small N-terminal helix, S1’s NTD makes an additional contact with the pRNA via a small loop (Fig. 5E). This second anchor point may cause S1’s position to deviate from helical symmetry.

(A) Side view of the ATPase NTDs showing their helical arrangement. The NTDs from five different subunits are labeled S1 to S5 and colored as in Fig. 4. ATP is colored by element (B) K56 is arranged as a spiral that approximately tracks the DNA helix. The bottom four subunits track the 5′-3′ strand approximately every 2 bp. Because of the imperfect NTD helical symmetry, the top K56 is closer to the complementary 3′-5′ strand. (C) Helical symmetry axes for successive superpositions of S1 on remaining subunits are shown as colored rods and labeled along with their actual and ideal (parentheses) rotations. Translational components of the helical operations are represented by the offsets of the rods. (D) COM of the NTD and CTD are shown as gray spheres from side and end-on views. The fivefold axis of the phage is shown as a darker gray rod. (E) S1 (blue) and its associated pRNA (magenta ribbon and translucent surface). The approximate portal-binding surface and the unique NTD-pRNA contact in S1 are shown as cyan and blue stars, respectively. (F) Superpositions of the pRNA from side and end-on views. The pRNAs were superimposed onto pRNA-S1 via their prohead-binding domains (bases 1 to 74) and colored according to their associated NTDs. The blue pRNA-S1 is oriented as in (E) note that S1 and S2 pRNAs are bent relative to the others such that its distal NTD-binding regions move up and to the right.

Since both the CTD and pRNA are arranged as planar rings, one question that arises is how the NTD can assume a helical arrangement when its anchor points are planar. While each NTD could bind to a different section of the pRNA to give rise to a staggered structure, our results suggest otherwise. Ignoring the unique third contact with pRNA made by S1 described above, all subunits essentially bind pRNA the same way. Instead, the pRNA changes conformation to adapt to the helical arrangement of NTDs (Fig. 5F). In this way, the pRNA functions analogously to the suspension of an off-road vehicle, where shock absorbers compress, extend, and flex to allow the wheels of the vehicle to maintain contact with the ground over uneven terrain. Similarly, the pRNA allows the NTD to maintain contact with its helical DNA substrate. However, unlike off-roader’s shock absorber, the A-form RNA helices cannot extend or compress, and thus, positional adjustments of the NTD are limited to bending and flexing motions. Thus, to “raise” the position of S1 to the top of the helix, the pRNA bends rather than simply shortening. As a result, in addition to positioning S1 closer to the portal, it is also slightly displaced in a direction approximately perpendicular to the fivefold axis of the phage. Since S1 is part of the larger NTD assembly, the whole cracked ring should be displaced in this same direction. The results presented here are consistent with this scenario. The pRNA attached to S1 is the most bent (Fig. 5E), as it must be to maintain S1’s position at the top of the helix. Further, plotting the centers of mass (COM) of the NTD and CTD show that, while the CTD COM lies on the global fivefold axis, the NTD COM is displaced as predicted (Fig. 5D). This explains why the DNA is bent since the DNA is nestled in the central pore of the NTD cracked ring, it is also displaced when the NTD cracked ring is displaced as it adopts a helical structure. Hence, there is likely a complex interplay between the NTD adopting a helical structure to bind DNA and the deformation of DNA structure in response to this binding.

Interactions with DNA

The central pores of both the CTD ring and the NTD helix of the ATPase are rich in positively charged and polar residues that are well positioned to interact with DNA (Figs. 3 to 5). Given the resolution of the map, it is difficult to definitively catalog these residues, but using a distance cutoff of 4 Å to select residues that could potentially interact with DNA shows that while the motor interacts with both strands, there seem to be more interactions with one of the two strands. This observation is consistent with single-molecule results showing that the motor tracks along the 5′-3′ strand during translocation (36). The overall helical character of the NTD described above is particularly apparent if one focuses on a single residue in the motor pore. Figure 5B shows K56, which appears to track along the 5′-3′ strand of the dsDNA helix approximately every 2 to 2.5 bp. Note that because of deviations from helical symmetry in both the dsDNA and the NTD cracked ring/helix, each K56 interacts with the DNA in a slightly different way. This is especially true for S1 at the top of the helix, which seems to be better positioned to interact with the opposite 3′-5′ strand, consistent with its deviation from strict C5 helical symmetry discussed above.

Intermolecular interactions between motor components

The N- and C-terminal domains are connected by a long linker analogous to the “lid” domains in other ASCE ATPases (Fig. 6). In addition to connecting the N- and C-terminal domains, the NTD-CTD linker also “links” adjacent subunits. It folds into a small three-helix domain, where the middle helix interacts with an adjacent subunit. Because of deviations in the helical pitch and twist, the orientation and the distance spanned by the linker arm are not uniform around the helix. The distance and orientation of the linker seem to be adjusted by varying the length of the two helices flanking the helix that binds the neighboring subunit more overall helical content shortens the distances, whereas direction can be altered by ending the helical region at different extents around a helical turn (Figs. 4C and 6C). The density for the linker regions from the subunits at the top (S1) and bottom (S5) of the ATPase helix is poor compared to the other three linker arms. This may be due to increased dynamics and/or conformational heterogeneity of these two linkers due to their unique positions in the NTD helix. In particular, the linker originating from the top subunit S1 must reach down to interact with the bottom subunit S5 and, therefore, must adopt a substantially different orientation compared to the other subunits. Subunit S5 at the bottom of the helix is also in a unique position and is part of the split interface between subunits S1 and S5. However, S5’s linker is not part of this interface but instead is part of the interface between subunits S5 and S4. Hence, the environment of S5’s linker should be similar to the environments of the three preceding subunits S4, S3, and S2. Nonetheless, similar to the top subunit, the bottom subunit’s overall position and orientation deviate from the stricter symmetry observed for the inner three subunits (Figs. 4C, 5C, and 6C). This difference in position and orientation may account for part of the increased flexibility observed for this region (see also below).

(A) Side view and (B) end-on view of adjacent subunits S1 and S2, colored as in Figs. 6 and 7. NTDs and CTDs are labeled, and the linker domain in S2 is highlighted in yellow. ATP is shown as space-filling spheres colored by element. (C) Superposition of the NTDs of all five subunits to illustrate structural variability of the linker domain. Note that the relative orientation of linker domains from subunits S1 (blue) and S5 (pink) differ the most and correspond to subunits at the top (S1) and bottom (S5) of the ATPase helix. (D) Close up of the active site between subunits S1 and S2. Residues important for binding and/or catalysis, including K105 and R146, are shown as ball and stick figures, colored by heteroatom, and labeled. ATP atoms are shown as ball and stick and colored by element note that phosphorous is colored light purple rather than the typical orange to facilitate visibility.

Active site of the ATPase

Although the resolution of the motor-vertex map is not sufficient to definitively visualize ATP, nucleotide binding sites of ASCE ATPases are well characterized, and the location of ATP can be inferred by superimposing the structure of a related ATPase complexed with ATP. In this case, superimposing the structure of the bacteriophage P74-26 DNA packaging ATPase complexed with substrate analog ADP-BeF3 (56) allowed positioning of substrate in active sites located between two adjacent subunits, consistent with previous data (31). In the middle three subunits (S2, S3, and S4), the nucleotide fits well into otherwise unexplainable density. However, in the top and bottom subunits (S1 and S5), there is no significant density corresponding to the expected position of the nucleotide. In addition, the linker arm closes over the active site and is thus positioned to respond to ATP binding, hydrolysis, and product release (Fig. 6, A and B). Density for the linker arm in S1 and S5 is poor, suggesting that nucleotide binding might influence the position and flexibility of the linker.

As described above, the active site resides between two subunits in the NTD helix. Thus, residues from two adjacent subunits contribute to a single active site, and each contributes to ATP binding and hydrolysis (Fig. 6D). In the “cis”-acting subunit, ATP is positioned near the Walker A and B motifs that reside on loops connecting strands on one edge of the central β sheet of the ASCE fold, consistent with previous results reported for phi29 and other ATPases. ATP is further stabilized by positively charged residues contributed in “trans” by an adjacent subunit. In particular, the ATPs seem to be sandwiched between R146 and K105 (Fig. 6D). In ring ATPases, an arginine residue often acts in trans to trigger ATP hydrolysis either by stabilizing the negative charge that accumulates on the phosphate oxygens during the hydrolytic transition state or by similarly stabilizing the negative charge on the ADP-leaving group. On the basis of sequence alignment with members of the FtsK branch of the ASCE superfamily, R146 was predicted to be the canonical ATPase “arginine finger.” This prediction was supported by biochemical experiments designed to report trans-acting activity (31, 35). While the current structural data could support R146 acting as the arginine finger, K105 seems equally well, if not better, positioned to assume this role. Specifically, K105 is positioned nearer the γ-phosphate of ATP (where nucleophilic attack occurs), whereas R146 is closer to the adenine base and pentose sugar (Fig. 6D). Further, an arginine residue equivalent to K105 in the thermophilic bacteriophage P74-26 was shown to function as the arginine finger (56). Hence, while R146 likely does act in trans during nucleotide cycling, assigning it the catalytic-trigger role of the arginine finger may be premature. Single-molecule optical laser tweezer experiments suggest that the R146 may play a role in nucleotide exchange (35), thus explaining the reported transactivity while allowing for the possibility that K105 could function as the catalytic “arginine” finger in gp16. Of note, in addition to closing over the active site, the linker interacts with the edge of the sheet in the trans-acting subunit where R146 and K105 reside (Fig. 6, B and D), consistent with potential roles in coordinating either/both ATP hydrolysis and nucleotide exchange in adjacent subunits.


X-Ray Crystallography Gives Scientists New Understanding Of Molecular Motor

Boston, MA (October 12, 1999) -- Harvard researchers have created the first atomic-resolution image of a donut-shaped enzyme, or helicase, that unwinds the DNA double helix to expose its genetic letters for DNA replication. Michael Sawaya, postdoctoral fellow in the lab of Tom Ellenberger, associate professor of biological chemistry and molecular pharmacology (BCMP), worked out the X-ray crystallographic structure reported in the October 15 Cell. The structure is rendered in pictures that show how six individual polypeptide lobes arrange themselves in space to look a bit like a ring of bread buns. It affords researchers the first detailed glance at a family of proteins that remain enigmatic in spite of their recognized status as a fundamental molecular machine of the cell.

"We know next to nothing about how these helicases move on DNA," says Ellenberger. Helicases have become a competitive field of inquiry because defects in human forms underlie several diseases, including the cancer-prone Bloom's syndrome and a disease of premature aging called Werner syndrome. Helicases are interesting for several reasons. First, they are molecular motors, just like myosin, which moves along actin fibrils to contract a muscle, or kinesin, which transports cargo along microtubules. The ring-shaped kind of helicase threads one strand of DNA through its central hole and zips along the double strand at breakneck speed, ploughing through 300 paired nucleotides per second while shoving the second strand out of the way. The enzyme is powerful, too. Other researchers have placed "roadblocks" in the helicase's way by binding proteins on the DNA's back. Yet the helicase knocked these off as it forced its way through.

Second, helicases came in different shapes--some are monomers, others dimers--and they do all sorts of things. The ring-shaped, hexameric helicase studied here spearheads a complex of enzymes as it pries apart the DNA strands for replication. Other helicases help with DNA repair, recombination, transcription, and more. They probably act wherever DNA needs to open up temporarily. Sequencing data suggests there are hundreds of different helicases in the human genome even the humble yeast boasts about 50 kinds.

For Ellenberger, the helicase represents a step in his ultimate goal to crystallize the entire replication fork of the bacteriophage T7, a complex of five different types of protein that copies DNA. Last year, researchers led by Ellenberger and Charles Richardson, professor of biochemistry and molecular pharmacology, reported the crystal structure of T7's DNA polymerase. Using biochemistry, Richardson's group had learned earlier that the helicase and the other proteins in the replication fork physically touch each other. Collectively, these interactions make the system work in still mysterious ways.

The crystal structure of the helicase does not, actually, represent the way this enzyme occurs in real life. In T7, the helicase is a double-decker protein with two enzyme activities. The large helicase donut sits atop a smaller one that is the primase, another enzyme of the replication fork. Richardson's lab prepared a helicase fragment of the helicase primase that was amenable to X-ray crystallography. Curiously, this fragment crystallized as an open ring, like a lock washer, whereas the biologically active form of the helicase more closely resembles a flat washer. "We think, however, that all interactions we are describing closely approximate what we would see in a closed ring," says Ellenberger.

So what did the scientists see? All amino-acid sites known to be conserved across helicases of this family from different species turned out to reside near the surface of the donut's hole, where one DNA single strand passes through. More importantly, though, the structure gives the researchers a first stab at solving the mechanism of how this motor generates motion from energy, like any engine does.

Scientists knew that the helicase splits a phosphate off dTTP--a relative of the fuel molecule ATP--to free up chemical energy. It must somehow convert this into the physical force needed to separate the Watson-Crick bonds joining the double-strand DNA base pairs. It also must harness energy for large changes in its shape, or conformation that allow it to step along the DNA. Finally, it needs a mechanism for grabbing and letting go of DNA.

The task lies in understanding precisely how the helicase's six subunits cooperate to make these things happen, and the crystal structure shows a plausible way, says Ellenberger. One dTTP is nestled in the cleft between every two, meaning that changes resulting from every dTTP reaction could spread to two subunits.

The dTTP binding site also abuts the DNA binding site at the donut's inner ring. When crystallized without dTTP, these DNA binding sites were disordered and consequently failed to appear as crisp patterns in the crystal structure. Yet when the researchers immersed the helicase crystals in a dTTP solution to allow the dTTP to seep into place, they found that not only was the dTTP sitting in its binding pocket but the adjoining DNA-binding region of the helicase also suddenly became visible. This suggests that the dTTP reaction might be coupled to DNA binding, in that dTTP cleavage enables the DNA binding region to "shape up" and grab the DNA. A fraction of a second later, this sequence could occur in the adjacent subunits, and so on throughout the ring.

Other researchers have advanced two major mechanistic models for simpler helicases that are now being hotly debated. Sawaya's structure cannot pick a winner largely because it is symmetrical. It does not visualize the larger conformational changes that must occur within the ring as it advances on the DNA.

The structure does, however, nurture an old passion, says Ellenberger. "As a kid I always loved engines, and I am still fascinated that you can make a protein function as this large, cooperative assembly to move rapidly down a strand of DNA. You've got all these motions flickering in a nanosecond time-realm. This works only because everything is supremely coordinated."

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Materials provided by Harvard Medical School. Note: Content may be edited for style and length.


What type of animation software is used to create these molecular motor motions? - Biology

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Cell Division

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Cell Biology and Cancer Animations (Rediscovering Biology)

How a Proto-oncogene Becomes an Oncogene: A depiction of some types of mutations that can occur to turn a proto-oncogene into an oncogene. p53's Role in the Cell: Shows various roles that p53 plays in the cell to protect the genome of the organism. Telomeres: Shows the concept of how the ends of chromosomes, the telomeres, shorten each time the cell divides. The Cell Cycle: Cyclins and Checkpoints: A depiction of the cell cycle and role that cyclins play in the process this animation also shows the role of checkpoints in regulating the cell cycle. The Signal Transduction Pathway: A depiction of the signal transduction pathway that is involved with the growth process of the cell.

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METHODS

Production of packaging components

Bacillus subtilis 12A (sup-) host cells were infected with the phi29 mutant phage sus 8.5(900)-16(300)-14(1241), which is defective in the packaging ATPase and thus cannot package DNA. The resulting empty prohead particles were purified via ultracentrifugation using sucrose gradients as previously described (68, 69). To ensure that the pRNA associated with these particles was structurally and compositionally homogeneous, purified particles were first treated with ribonuclease A to remove any attached pRNA and repurified as previously described (69). In parallel, 120 base pRNA was produced from the plasmid pRT71 by in vitro transcription using T7 RNA polymerase and purified by denaturing urea polyacrylamide gel electrophoresis as previously described (70). The RNA-free proheads were then reconstituted with uniform 120 base as previously described (69). The genomic DNA-gp3 substrate and packaging ATPase gp16 were prepared and purified as previously described (31, 68).

DNA packaging assay

The in vitro DNA packaging assay is based on a deoxyribonuclease (DNase) protection assay and was performed as previously described (68, 69). Briefly, proheads (8.3 nM), DNA-gp3 (4.2 nM), and either wild-type or mutant gp16 (166 to 208 nM) were mixed together in 0.5× tris(hydroxymethyl)aminomethane-HCl magnesium salt (TMS) buffer in 20 μl and incubated for 5 min at room temperature. ATP was then added to 0.5 mM to initiate packaging. After 15-min incubation, the mixture was treated with DNase I (1 μg/ml final concentration) and incubated for 10 min to digest the unpackaged DNA. An EDTA/proteinase K mix was then added to the reaction mixture (25 mM and 500 μg/ml final concentration, respectively) and incubated for 30 min at 65°C to inactivate the DNase I and release the protected, packaged DNA from particles. The packaged DNA was analyzed by agarose gel electrophoresis. Packaging efficiency was calculated by densitometry using the UVP Gel Documentation System.

ATPase assays

ATPase activity of prohead/gp16 (wild-type or mutant) motor complexes was determined by measuring production of inorganic phosphate by the EnzChek Phosphate Assay Kit (Life Technologies) as described previously (43). Briefly, a reaction mixture containing reaction buffer 2, and 0.1 mM sodium azide] or Tris-HCl magnesium (TM) buffer [25 mM tris (pH 7.6) and 5 mM MgCl2]> and 0.2 mM 2-amino-6-mercapto-7-methylpurine riboside) with proheads (4.2 nM) and gp16 (125 nM) in 90 μl was preincubated at room temperature for 10 min in the presence of purine nucleoside phosphorylase (0.1 U). ATP was added to 1 mM to initiate the reaction and production of Pi measured in the spectrophotometer at A360 for 10 min.

Production of DNA packaging intermediates for cryo-EM

To generate the stalled DNA packaging intermediates for cryo-EM reconstruction, packaging reactions consisting of DNA-gp3, RNA-free proheads reconstituted with 120 base pRNA and gp16 were assembled at 2× concentration (see above), and ATP was added to initiate DNA packaging. Three minutes after ATP addition, ATP-γ-S was added to 100 μM (Roche) (ATP concentration is 500 μM) and incubated for 2 min. To maximize packaging efficiency, a prohead-to-DNA ratio of 2:1 was used. Hence, at most, only 50% of the particles could package DNA. However, far fewer particles had actually packaged DNA (Fig. 2), and thus, there was considerable amounts of unpackaged DNA fibers in the background of cryo-EM images. Thus, to facilitate freezing and remove background noise contributed by unpackaged DNA, 1 U of RQ DNase I (Promega) was added and incubated at room temperature for 10 min. The sample was placed on ice until grid preparation for cryo-EM imaging.

While enough gp16 was added to assemble functional motors on every particle, the large number of unpackaged particles raised concerns that motors on these particles were either not fully or properly assembled. Thus, the reaction mixture was loaded on a sucrose gradient (with an excess of ATP-γ-S included) to separate DNA-filled particles from empty particles. The gradient band corresponding to filled particles was dialyzed against TMS buffer supplemented with ATP-γ-S to remove sucrose, concentrated, and loaded onto cryo-EM grids for freezing and imaging. The resulting grids had a good distribution of almost entirely filled particles. Since most particles in the images had packaged DNA, we could presume that their packaging motors were functional. Although a small amount of “unpackaged” DNA was still present, this likely resulted from some particles losing their DNA during purification and was not enough to hinder image processing. Although these empty particles had likely packaged but then lost DNA, they were nonetheless excluded from image processing to ensure that only functional motors in the same state were reconstructed.

Cryo-EM grid preparation, data collection, image processing, and model building

Approximately 3 μl of prohead particles stalled during packaging (see above production of packaging intermediates for cryo-EM) was applied to quantifoil 2 × 2 holey carbon grids before plunge-freezing. Sample grids were flash-frozen in liquid ethane cooled to liquid nitrogen temperatures on holey carbon grids using a vitrobot automated vitrification system from Thermo Fisher Scientific. Data were collected on the Titan Krios microscope equipped with a summit K2 direct electron detector housed at the Electron Imaging Center for Nanomachines at University of California, Los Angeles. The accumulated electron dosage for each movie stack was

40 e/Å 2 . Subsequent Contrast transfer function (CTF) correction (see below) indicated a defocus range of

3.5 μM. Individual movie frames were aligned with Motioncor2 (71), and particles were picked with EMAN2 (72). Subsequent image processing steps, including symmetry breaking and focused reconstruction, were carried using Relion (73) and/or symmetry breaking scripts written for Relion (39) and as implemented in Scipion (74) (Supplementary Materials). For the DNA-filled particles discussed here, a total of 145,434 particles were initially boxed and extracted (fig. S1). After removing incorrectly picked or deformed particles via 2D classification, 53,010 particles were used for the reconstruction of the entire phage. For the focused reconstruction including the ATPase motor, only particles where motors were clearly visible were used, reducing the number of particles to 12,526. Atomic models of the motor components were fitted and/or manually built into their corresponding densities using COOT (75) and refined using PHENIX (76). In the final refined model, 90.28% amino acid residues were in the preferred region of the Ramachandran plot, with no residues in the disallowed region and 9.72% in the generously allowed region. The MolProbity score of the model was 2.45. The correlation coefficient between density calculated from the refined structure and the cryo-EM map of the motor vertex was

0.63. Both the FSC between the model, the map, and the d_model (76) indicated resolutions of

4.33 Å for the masked map. For the unmasked map, the d_model resolution estimate was

4.5 Å (fig. S1), whereas the FSC estimate was

6.19 Å, likely due to unmodeled DNA and residual capsid protein in the unmasked map. The focused reconstruction of the motor vertex and associated atomic model of the pRNA, ATPase, and DNA are deposited as EMD-22441 and PDB ID: 7JQQ, respectively.